by...to my lab-mates over the years, dr melvin leteane, dr azimah, david, salmah, nargs, mariam,...
TRANSCRIPT
INVESTIGATING RNA GRANULES
FORMATION DURING
CALICIVIRUSES INFECTION
By
MAJID NOORI HUMOUD AL-SAILAWI
THESIS SUBMITTED FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY
FACULTY OF HEALTH AND MEDICAL
SCIENCES
UNIVERSITY OF SURREY
May 2015
i
ABSTRACT
Human norovirus (HuNV) is a member of the calicivirus family and is a major cause of
viral gastroenteritis worldwide. Due to the absence of a suitable cell culture system, HuNoV
replication mechanisms are poorly understood, but two animal caliciviruses, Feline
calicivirus (FCV) and Murine Norovirus (MNV) provide models to increase our
understanding of norovirus biology. Unlike cellular mRNAs, the calicivirus RNA genome
does not possess a 5' cap structure but instead has a 13–15 kDa viral protein, genome linked
(VPg) directing translation, hijacking the host protein synthesis machinery. The viral life
cycle requires separated events occurring at different times since viral transcripts are used as
the template both for translation (mRNA) and replication (genomic RNA). Therefore
mechanisms are required to control the viral RNA fate. In eukaryotes, during stress
conditions, mRNAs can be stored in subcellular compartments such as stress granules to stall
their translation or in processing bodies to be degraded. Recent evidence indicates that these
compartments also play an important role during the viral life cycle. Therefore, using
immunofluorescence microscopy we set out to investigate how FCV and MNV infection
regulate the formation of G3BP1- and PABP-1-containing stress granules and DCP-1-
containing processing bodies to address whether these cytoplasmic granules could play a role
during the viral life cycle. We have now shown that FCV has the ability to prevent stress
granules formation during infection and that this is important for replication in CRFK and
FEA cells. Using FCV-free supernatant from infected CRFK cells and immunofluorescence
microscopy, we have also shown that during infection, the formation of stress granules is
induced in a paracrine manner in uninfected cells via a messenger molecule released from
infected cells. We hypothesize that this could reflect a new antiviral role for stress granules.
ii
Furthermore, MNV and FCV infection also led to the disruption of processing-bodies
assembly. Overall, this study revealed that caliciviruses modulate the RNA granules during
infection and that this could be part of viral mechanism to counteract the antiviral response.
iii
ACKNOWLEDGEMENTS
I would like to say a huge thank you to my supervisor Dr Nicolas Locker for his
advice, endless patience, guidance, inspiration and support. It has been pleasure to working
under his supervision and I am lucky to have worked with him. Without his supervision and
constant help my dissertation would not have been possible.
I would also like to thank my co-supervisor Professor Lisa Roberts for her guidance,
encouragement and support. I am extremely grateful to her.
A massive thank you also goes to Dr Elizabeth Royall for her assistance, support, proof
reading and brilliant comments and suggestions. I also need to thank Dr Nicole Doyle and Dr
Margaret Carter. Thank you for all your help, support and your friendship. I would also like
to thank Dr Dominique Weill in Université Paris 6 (Paris, France) for her advice. I would also
like to thank Dr. Belinda Hall for her advice. Thanks also to Dr Rachel Butler and Robert
Francis for their help and advice with the confocal work.
To my lab-mates over the years, Dr Melvin Leteane, Dr Azimah, David, Salmah, Nargs,
Mariam, Nefeli and Hanan, a big thank you for the all fun times, your friendship and support.
Thank you to the University of Basra for proving funding for my study.
My heartfelt thanks to my extended family and my wife family for their encouragement,
prayers and helping me over these four years.
To soul of my father and my granny, thank you to painting me in the right way. I miss you.
Finally to my mother, brother and his family, my sister, my wife and my children Mohammed
Ridha and Yusuf. You have lived through all the ups and downs of this PhD with me, and for
have given me your constant love, support and encouragement. I could not have done this
without you (no thank you can ever be enough).
It’s all for you.
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TABLE OF CONTENT
CHAPTER 1 Introduction 1
1.1 RNA Granules 2
1.2 Cytoplasmic mRNPs granules (Stress granules and P-bodies) 3
1.3 Stress granules (SGs) 4
1.3.1 Stress granule composition 6
1.3.2 Stress granule assembly/disassembly 11
1.3.3 Functions and importance of stress granules 17
1.4 Processing bodies (P-bodies or PBs) 20
1.4.1 P-body assembly/disassembly 23
1.4.2 Processing bodies functions 25
1.4.3 Role of PBs in mRNA degradation 26
1.4.4 Role of P-bodies in mRNA surveillance 28
1.4.5 Role of P-bodies in gene silencing 28
1.4.6 Other functions of PBs 29
1.5 Interactions between Stress granules and P-bodies 30
1.6 Stress granules, P-bodies, and viral life cycles 35
1.6.1 Viral interaction with stress granules 36
1.6.2 Inhibition of Stress granules by viral proteinase 38
1.6.3 Inhibition of stress granule formation by modulation of eIF2α phosphorylation 39
1.6.4 Inhibition of stress granule formation by sequestering or co-opting their
components
40
1.6.5 Viral replication mechanism that benefits from SG formation 43
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1.7 Virus/P-bodies interaction 45
1.8 Caliciviruses 48
1.8.1 Lagoviruses (LaVs) 51
1.8.2 Vesiviruses (VeVs) 52
1.8.3 Sapoviruses (SaVs) 53
1.8.4 Neboviruses (NeVs) 55
1.9 Noroviruses (NoVs) 55
1.9.1 Animal noroviruses 57
1.9.2 Murine norovirus-1 (MNV-1) 57
1.9.3 Human noroviruses (HuNoVs) 58
1.9.4 Clinical signs 61
1.9.5 Transmission 62
1.10 Model systems for the study of human norovirus biology 63
1.11 Pathogenesis and immunity 67
1.12 Viral structure, genome organisation and function of the viral proteins 70
1.13 Norovirus life cycle 78
1.14 The function of RNA-protein interaction during noroviruses infections 84
1.15 Aims and objectives of the research 88
CHAPTER 2 General Materials and Methods 89
2.1 Maintenance of cells 90
2.1.1 Crandall Rees Feline Kidney (CRFK) Cells 90
2.1.2 Mouse Leukemic monocyte-macrophage Cells (RAW 264.7) 90
2.1.3 Feline embryonic airway (FEA) cells 91
2.1.4 Murine microglial cells (BV-2) 91
vi
2.1.5 Human embryonic kidney 293 (HEK293T) cells (293 cells) 92
2.1.6 Murine macrophage J774 cells 92
2.1.7 THP-1 cells 93
2.2 Freezing and Resuscitation of CRFK and RAW264.7 cells 93
2.3 Preparation of viruses stock 94
2.3.1 Preparation of Feline Calicivirus (FCV) stock 94
2.3.2 Production of Murine Norovirus (MNV-1) stock 94
2.4 50% Tissue Culture Infectious Dose (TCID50) Assay 95
2.5 Preparation of VPg-linked FCV RNA 95
2.6 Preparation of VPg-linked MNV RNA 96
2.7 Preparation and quantitation of VPg-linked FCV and MNV RNA 97
2.8 Immunofluorescence studies 97
2.8.1 Cell culture and virus infection 97
2.8.2 Immunofluorescence assay 97
2.9 Effect of calicivirus infection on SGs and PBs formation 98
2.9.1 Staining for FCV-p76 protein 100
2.9.2 Staining for MNV-3D polymerase protein 100
2.9.3 Staining for SG-markers 100
2.9.4 Staining for PBs-markers 101
2.10 SGs formation during FCV infection 101
2.10.1 Treatment with sodium arsenite (NaAsO2) 101
2.10.2 Effect of FCV infection on SGs formation 102
2.10.3 Effect of NaAsO2–induced SGs on FCV replication 102
2.10.4 Effect of FCV infection on hydrogen peroxide (H2O2) induced SGs 102
2.10.5 Effect of the H2O2-induced SGs on FCV replication 103
vii
2.11 Preparation of FCV free supernatants 103
2.11.1 Virus infection and clarification (virus elimination) 103
2.11.2 Virus Precipitation 105
2.13 Virus inactivation using ultraviolet-light (UV-light) 105
2.14 Effect of cell culture supernatant on stress granules formation 106
2.15 Effect of interferon on stress granules formation 106
2.16 Effect of RNase A treatment on stress granules formation 106
2.16 Effect of heat shock on stress granules formation 107
2.17 SGs formation during MNV-1 infection 107
2.18 Formation of P-bodies during FCV Infection. 108
2.19 Formation of P-bodies during MNV-1 Infection 108
2.20 Statistical analysis
108
CHAPTER 3 Regulation of Stress Granules formation during FCV and MNV-1
infection
110
3.1 Optimisation of experimental conditions for infection and SGs detection 111
3.2 Chemical induction of SGs using oxidative stress in CRFK cells 117
3.3 Optimisation of the markers and conditions used for stress granules detection
in CRFK cells
119
3.4 Optimisation of the markers used for stress granules detection in RAW264.7
cells
123
3.5 Stress granules formation during FCV infection. 125
3.6 Stress granules formation during MNV-1 infection. 134
3.7 FCV infection impairs the assembly of stress granules induced by sodium
arsenite
139
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3.8 FCV infection impairs the assembly of stress granules induced by hydrogen
peroxide
145
3.9 Effect of Stress granules induction on FCV replication 149
3.10 Effect of FCV infection on pre-assembled stress granules 151
3.11 Effect of UV-inactivation on the inhibition of stress granules assembly 156
3.12 Discussion 160
3.12.1 Absence of stress granules in stressed and MNV-infected RAW264.7
macrophages
161
3.12.2 FCV replication impairs stress granules assembly during infection
162
CHAPTER 4 Paracrine induction of Stress Granules by FCV-infected cells 167
4.1 Generation of virus-free cell culture supernatant 168
4.2 Effect of FCV-infected cells supernatant on Stress granule formation 171
4.3 Effect of interferon α on stress granule formation in CRFK cells 174
4.4 Effect of Ribonuclease A (RNase A) on the ability of FCV-infected cells
supernatant to induce stress granule assembly
178
4.5 Effect of Heat-Shock on the ability of FCV-infected cells supernatant to
induce stress granule assembly
181
4.6 Effect of FCV-infected cells supernatant on wide range of cells’ ability to
assemble stress granules
185
4.7 Discussion 188
4.7.1 FCV infection leads to paracrine induction of SG assembly 188
4.7.2 Different types of stress granules can be assembled in response to infection 189
4.7.3 Could this paracrine induction of SGs reflect antiviral mechanism? 190
4.7.4 Investigating the nature of the soluble mediator(s) triggering stress granules 192
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assembly
CHAPTER 5 Regulation of P-Bodies formation during FCV and MNV-1 infection
5.1 Optimisation of the markers and conditions used for P-bodies detection 195
5.2 Effect of MNV-1 infection on P-bodies formation 200
5.3 Effect of FCV infection on the assembly of P-bodies 210
5.4 Effect of chemical induction of oxidative stress on P-bodies assembly 213
5.5 Discussion 217
5.5.1 Modulation of P-bodies assembly during MNV-1 and FCV infection 217
6.1 CHAPTER 6 Conclusion 223
7.1 CHAPTER 7 Reference 229
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LIST OF FIGURES
CHAPTER 1
1.1 Proposed model of induction stress granules (SG) assembly by different
conditions and stimuli
13
1.3 Eukaryotic mRNA decay pathways 27
1.3 Components of P-bodies and stress granules 31
1.4 Proposed model for the role of cytoplasmic RNA granules in mRNA fate 34
1.5 Mechanisms of stress granules disruption by viruses 37
1.6 Mechanisms of P-bodies disruption by viruses 47
1.7 Crystal structures of Norwalk Virus, MNV-1 and FCV by cryo-electron
microscopy
50
1.8 Phylogenetic analysis of the capsid amino acid sequence of Norovirus strains 56
1.9 Organization of representative caliciviruses genome 72
1.10 The structure of Norovirus major capsid protein, viral protein 1 (VP1) 77
1.11 Proposed replication mechanism of the Human Norovirus (HuNoV) 79
1.13 Schematic diagram of cap and VPg-dependent translation initiation
82
CHAPTER 2
2.1 Preparation of the FCV-free supernatant from FCV-infected cells for
immunofluorescence studies
104
CHAPTER 3
3.1 Optimization of FCV MOI for immunofluorescence studies 113
3.2 Optimization of MNV-1 MOI for immunofluorescence studies 115
3.3 Optimization of p76 antibody dilution for immunofluorescence studies 116
3.4 Induction of stress granules by oxidative stress 118
3.5 Detection of stress granules by oxidative stress in FCV-infected cells using
various markers
120
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3.6 3.6 Optimization of FBS concentration for detect of stress granules in CRFK
cells
122
3.7 Detection of stress granules in RAW264.7 cells using various markers 124
3.8 Detection of stress granules during FCV infection 126
3.9 Quantification of the number of cells displaying stress granules during FCV
infection
127
3.10 Detection of stress granules during FCV infection using PABP-1 as a marker 129
3.11 Quantification of the number of cells displaying stress granules during FCV
infection using PABP-1
130
3.12 Detection of SGs in FEA cells during FCV infection 131
3.13 Detection of SGs formation during FCV infection using different markers 133
3.14 Detection of stress granules during MNV-1 infection 135
3.15 Detection of stress granules during MNV-1 infection using various markers 136
3.16 Detection of stress granules during MNV-1 infection in BV-2 cells using
different markers
138
3.17 Detection of stress granules induced by NaAsO2 following FCV infection 140
3.18 Quantification of the number of cells displaying stress granules induced by
NaAsO2 following FCV infection at different time points
141
3.19 Detection of stress granules induced by sodium arsenate (SA) following FCV
infection using PABP-1 as a marker
143
3.20 Quantification of the number of cells displaying stress granules induced by
sodium arsenate following FCV infection using PABP-1 as a marker
144
3.21 Detection of stress granules induced by hydrogen peroxide (H2O2) following
FCV infection
147
3.22 Quantification of the number of cells displaying stress granules induced by 148
xii
hydrogen peroxide (H2O2)
3.23 Effect of stress granules induction on FCV replication 150
3.24 Effect of FCV infection on pre-assembled stress granules 154
3.25 Quantification of the number of cells displaying stress granules upon sodium
arsenate (SA) treatment and FCV infection
155
3.26 Effect of UV-inactivation on FCV replication 157
3.27 Effect of UV-inactivation on the inhibition of Stress Granules assembly 158
3.23 Quantification of the number of cells displaying stress granules during FCV or
UV-inactivated FCV infection and sodium arsenate (SA) treatment
159
CHAPTER 4
4.1 Detect of FCV replication by TCID50 following virus elimination 170
4.2 Induction of stress granules by cell supernatant 172
4.3 Quantification of the number of cells displaying stress granules following
treatment with cell supernatant
173
4.4 Effect of interferon treatment on stress granule formation 176
4.5 Quantification of the number of cells displaying stress granules following
treatment with IFNα
177
4.6 Effect of RNase A treatment on stress granule formation 179
4.7 Quantification of the number of cells displaying stress granules following
treatment with RNase A
180
4.8 Effect of heat shock on stress granule formation 183
4.9 Quantification of the number of cells displaying stress granules following
treatment with heat shocked supernatant
184
4.10 Effect of FCV-infected cells supernatant on wide range of cells 186
4.11 Quantification of the number of cells displaying stress granules following 187
xiii
treatment wide range of cells with FCV-infected cell supernatant
CHAPTER 5
5.1 Optimization of Dcp1 and Xrn1 antibody dilution in CRFK cells for
immunofluorescence studies
196
5.2 Optimization of Dcp1 and Xrn1 antibody dilution in RAW264.7 cells for
immunofluorescence studies
197
5.3 Optimization of foetal bovine serum concentration for detect of P-bodies in
CRFK cells
199
5.4 Detection of P-bodies during MNV-1 infection 202
5.5 Quantification of the number of cells displaying P-bodies during MNV-1
infection
203
5.6 Detection of P-bodies during MNV-1 infection using BV-2 cells 205
5.7 Quantification of the number of cells displaying P-bodies during MNV-1
infection in BV-2 cells
206
5.8 Detection of P-bodies during MNV-1 infection using Xrn1 as a marker 209
5.9 Detection of P-bodies during FCV infection 211
5.10 Quantification of the number of cells displaying P-bodies during FCV infection 212
5.11 Effect of the sodium arsenite and hydrogen peroxide treatment on P-body
assembly
215
5.12 Quantification of the number of cells displaying P-bodies upon treatment with
the sodium arsenite and hydrogen peroxide
216
CHAPTER 6
6.1 Proposed working model for the effect of FCV and MNV-1 on stress granules
and P-bodies.
228
xiv
LIST OF TABLES
CHAPTER 1
1.1 Components of stress granules 8
1.2 Components of P-bodies 22
1.3 Taxonomy of the Caliciviridae family
49
CHAPTER 2
2.1 Primary and secondary antibodies that are used in immunofluorescence studies 99
xv
LIST OF ABBREVIATIONS
2Apro 2Apro protease
3Cpro 3C-like Cysteine protease
4E-BP eIF4E-binding protein
40S Eukaryotic small ribosomal subunit (40 Svedberg)
60S Eukaryotic larg ribosomal subunit (60 Svedberg)
APOBEC3G Apolipoprotein B mRNA-editing enzyme, catalytic polypeptide-like 3G
APOBEC3F Apolipoprotein B mRNA-editing enzyme, catalytic polypeptide-like 3F
ADAR Adenosine deaminase, RNA-specific
Ago2 Argonaute RISC catalytic component 2
ALS amyotrophic lateral sclerosis
ATF4 Activating transcription factor 4
ATP Adenosine triphosphate
BNoV Bovine norovirus
BRF1 Butyrate response factor 1
Caprin-1 Cell cycle associated protein 1
CRFK Crandell Rees Feline Kidney cells
CIRP Cold-inducible RNA-binding protein
CPEB Cytoplasmic polyadenylation element binding protein
CrPV Cricket paralysis virus
CVB3 coxsackievirus B3
GADD34 DNA damage–inducible protein 34
DCP1 Decapping mRNA 1
DDX3 DEAD box helicase 3
DDX6/RCK DEAD box helicase 6
xvi
DMEM Dulbecco’s Minimum Essential Medium
DMSO Dimethyl sulfoxide
DV Dengue virus
EBHSV European Brown Hare Syndrome Virus
eEF2 eukaryotic elongation factor 2
Edc1-3 Enhancer of decapping protein 1-3
EDC4/GE-1/Hedls Enhancer of mRNA decapping 4
EMCV encephalomyocarditis virus
eIF Eukaryotic initiation factor
eIF1A Eukaryotic translation initiation factor 1A
eIF2 Eukaryotic translation initiation factor 2
eIF3 Eukaryotic translation initiation factor 3
eIF4A Eukaryotic translation initiation factor 4A
eIF4B Eukaryotic translation initiation factor 4B
eIF4G Eukaryotic translation initiation factor 4G
eIF4H Eukaryotic translation initiation factor 4H
eIF5B Eukaryotic translation initiation factor 5B
EMCV Encephalomyocarditis virus
EV71 Enterovirus 71
FAST Fas-activated Ser/Thr kinase
FBS Fœtal Bovine Serum
FCV Feline calcivirus
FCV-VSD FCV-associated virulent systemic disease
FEA Feline Embryonic Airway
FMDV Foot-and-mouth disease virus
xvii
FMR Fragile X mental retardation
FAK focal adhesion kinase
FRMP Fragile X mental retardation protein
FUS Fused in Sarcoma
FXS Fragile X syndrome
O-GalNAc O-N-acetylgalactosamine
G3BP1 Ras-GTPase-activating protein SH3-domain-binding protein 1
GCN2 General control non-derepressible 2
GDP Guanosine diphosphate
gRNA Genomic RNA
Grb7 growth factor receptor-bound protein 7
GTP Guanosine triphosphate
GW182/ TNRC6A GWbodies 182/ Trinucleotide repeat-containing 6A
(HBGAs) Histo-blood group antigens
HCV Hepatitis C virus
HDAC6 Histone deacetylase 6
HIV-1 human immunodeficiency virus 1
HLTV-1 human T-cell leukaemia virus type 1
HRI Heme-regulated eIF2a kinase
HIV Human Immunodeficiency Virus
hpi Hour post infection
HSGs Heat shock granules
HSP Heat shock protein
HSV-1or 2 herpes simplex virus-1 or 2
HuNoV Human Norovirus
xviii
HuR/ELAVL1 Hu antigen R/ELAV-like RNA-binding protein 1
IAV influenza A virus
ICTV International Committee on Taxonomy of Viruses
IP5K Ins Intracellular localization of human Ins(1,3,4,5,6)P5 2-kinase
IFN Interferon
JAM-A Junctional Adhesion Molecule A
JEV Japanese encephalitis virus
JUNV Junín virus
KSHV Kaposi’s Sarcoma-associated Herpesvirus
kDa Kilo Dalton
LaVs Lagoviruses
MBNL1 Muscleblind-like protein 1
MCPIP1 Monocyte chemotactic protein-induced protein 1
MEF Mouse embryo fibroblasts
MEM Eagle’s Minimum Essential Medium
(MHE) Mouse hepatitis coronavirus
miRNA Micro RNA
MLN51 Metastatic lymph node 51
MNV Murine Norovirus
MOI Multiplicity of Infection
mRNA messenger RNA
mRNPs messenger ribonucleoproteins
MRV mammalian orthoreovirus
mTOR Mammalian target of rapomycin
M7GpppG 7-methyl guanosine cap structure of RNA
xix
NeVs Neboviruses
NHS UK National Health Service
NMD Nonsense-mediated decay
NoVs Noroviruses
NS Non-structural protein
nt nucleotide
NV Norwalk Virus
ORF Open reading frame
PABP Poly (A)-Binding Protein
Pat A Pateamine A
PCBP2 Poly (rC)-binding protein
PKR Protein kinase R
PEC porcine enteric calicivirus
PERK Protein kinase RNA-like endoplasmic reticulum kinase
PSaV Porcine SaV
PV Poliovirus
PIC Pre-initiation complex
Poly (A) Polyadenylation tail
PTB Polypyrimidine tract-binding protein
RACK1 The Receptor for Activated C Kinase 1
RAG2 Mice lacking recombination-activation gene 2
RAP55/LSM14A RNA-associated protein 55
RdRp RNA dependent RNA polymerase
RHDV Rabbit Haemorrhagic Disease Virus
RISC RNA-induced silencing complexes
xx
Rpb4 RNA polymerase II subunit B4
RRL Rabbit Reticulocyte Lysate
RSV Respiratory syncytial virus
RT Room Temperature
rVLPs Recombinant virus like-particles
SaVs Sapoviruses
SDS-PAGE Sodium dodecyl sulphate polyacrylamide gel electrophoresis
SFV Semliki Forest Virus
sgRNA Sub-genomic RNA
siRNA Small Interfering RNA
SMN Survival of motor neuron
SMSLV San Miguel Sea Lion Virus
STAT 1 Signal Transducers and Activators of Transcription
TDP-43 TAR DNA-binding protein 43
TCID50 50% Tissue Culture Infectious Dose
TC Ternary complex
TH2RNP TIAR-HIV-2 ribonucleoprotein
TIA1 T cell restricted intracellular antigen-1
TIAR TIA-1-related protein
TMEV Theiler’s murine encephalomyelitis virus
TRAF2 TNF receptor-associated factor 2
UPF1,2,3 Regulator of nonsense transcripts 1,2,3
USP10 Ubiquitin specific peptidase 10
TTP Tristetraprolin
TV Tulane virus
xxi
UTR Untranslated region
UV Ultra-violet
VeVs Vesiviruses
VLP Virus-Like Particle
VP Virion Protein
VPg Viral Protein Genome-linked
VSV Vesicular Stomatitis Virus
VESV Vesicular Exanthema of Swine Virus
vhs Viral host shutoff protein
wt Wild Type
WHO Word Health Organisation
WNV West Nile virus
Xrn1 5’-3’ Exoribonuclease 1
ZBP1 Z-DNA-binding protein 1
1
Chapter 1
Introduction
2
1.1 RNA Granules
RNA granules are self-assembling structures, or foci, that form in response to stress
conditions via the aggregation of non-translating mRNAs with several proteins to form
messenger ribonucleoproteins (mRNPs) granules (Anderson & Kedersha, 2006; Buchan,
2014; Reineke & Lloyd, 2013; Tsai & Lloyd, 2014). The assembly of these membrane-less
granules occurs either in the nucleus (nuclear mRNPs granules) or in the cytoplasm
(cytoplasmic mRNPs granules) and both are found in all type of eukaryotic cells, contributing
to many events of mRNA metabolism and processing (such as translation, subcellular
localization, turnover) (Balagopal & Parker, 2009; Buchan, 2014; Reineke & Lloyd, 2013).
Nuclear granules including Cajal bodies, nuclear speckles, Polycomb bodies,
paraspeckles, histone locus bodies and nuclear stress bodies are involved in the modulation of
several nuclear activities such as RNA splicing, modification, assembly and storage of small
nuclear RNP (snRNPs), mRNA maturation and export, histone mRNA transcription and
processing, gene transcription and others (Anderson & Kedersha, 2009a; Caudron-Herger &
Rippe, 2012; Mao et al., 2011). The cytoplasmic RNA granules are classified according to
cellular context, marker proteins, and function into various types including stress granules
(SGs), Processing-bodies (P-bodies; PBs) in somatic cells, germ granules in germ cells and
neuronal granules in neurons (Buchan, 2014). These granules have been implicated in innate
immunity, post-transcriptional regulation of gene expression, mRNA storage and mRNA
decay or degradation (Anderson & Kedersha, 2006).
Germ granules of various classes are found in germ cells dependent on their
development stage including nuage (germ cell), sponge, balbiani, and chromatid bodies
(gametes) and germ plasma (embryos). They contain several proteins such as RNA helicase
Vasa involved during germ cell development events such as regulation, localisation, stability,
3
translation and storage of mRNAs (Voronina et al., 2011). In addition, some proteins
involved in mRNA translation such as eukaryotic initiation factor 4E (eIF4E) (Amiri et al.,
2001), and RCK (Navarro & Blackwell, 2005), accumulate in germ granules, as well as
proteins involved in mRNA degradation including Sm/Lsm proteins (Audhya et al., 2005;
Boag et al., 2005), and Decapping proteins 1 and 2 (Dcp1/2) (Lall et al., 2005).
Neuronal RNA granules, also termed transport granules, are mainly involved in
mRNA transport but also play a possible role in mRNA localisation (Anderson & Kedersha,
2006; Krichevsky & Kosik, 2001). In addition to proteins such as Staufen (Thomas et al.,
2005), G3BP1 and HuR/D (Atlas et al., 2004), these granules contain ribosomal subunits
(40S and 60S) and translation initiation factors (eIF2, eIF3 and eIF4E) (Kiebler & Bassell,
2006; Krichevsky & Kosik, 2001).
The most common cytoplasmic granules associated with mRNA turnover are SGs and
PBs. SGs contain non-translating mRNA along with 40S ribosomal subunits as well as a
subset of translation initiation factors and RNA binding proteins, while PBs are enriched with
proteins involved in mRNA degradation and decay (Jain & Parker, 2013a). All these types of
RNA granules contain non-translated mRNA and share many RNA binding proteins which
may reflect the possible relationship between these structures (Buchan, 2014).
1.2 Cytoplasmic mRNPs granules (Stress granules and P-bodies)
In order to maintain cellular homeostasis during changing conditions caused, for
example, by stress, somatic cells apply a variety of regulatory mechanisms allowing the
control of gene expression. One of these mechanisms is the global inhibition or
reprogramming of mRNA translation in response to stress conditions; this impairs gene
4
expression and contributes to mRNA turnover until stress recovery (Shenton et al., 2006).
Cytoplasmic SGs and PBs form in response to translational arrest via the aggregation of non-
translated mRNAs with a variety of proteins, they play critical roles in regulating gene
expression via the control of mRNA translation and degradation during stressful conditions
such as viral infections (Anderson & Kedersha, 2002; Brengues et al., 2005; Decker &
Parker, 2012; Parker & Sheth, 2007). In eukaryotic cells, cytosolic mRNAs are in dynamic
equilibrium among polysomes, where mRNAs are translated, SGs, in which untranslated
mRNAs and associated proteins are stored during conditions of stress, and PBs, which
contain the decay machinery for degrading non-translated mRNA (Anderson & Kedersha,
2006; Parker & Sheth, 2007). The movement of mRNA between these different functional
states may support the roles of these subcellular structures in regulating gene expression
(Brengues et al., 2005; Buchan et al., 2008; Decker & Parker, 2012; Kedersha et al., 2005;
Mollet et al., 2008). Despite the differences between PBs and SGs in composition and
function, both are induced by various types of environmental stress such as heat shock, UV
light, starvation, oxidative stress (arsenite poisoning), and endoplasmic reticulum stress, as
well as under certain physiological conditions and during viral infections (Kedersha et al.,
1999; Towers et al., 2011; Tsai & Lloyd, 2014). Furthermore, they share many components
as detailed in the next section (Buchan, 2014; Kedersha & Anderson, 2009).
1.3 Stress granules (SGs)
SGs were discovered in 1983 as foci, or spherical granules, containing a specific
subset of cellular mRNAs within the cytoplasm of Peruvian tomato (Lysopersicon
peruvianum) cells subjected to heat shock in vitro (Nover et al., 1983). The formation of
these foci was associated with the inhibition of translation in stressed tomato cells and these
5
granules termed heat shock granules (HSGs), containing a 27-kDa heat shock protein
(HSP27) as well as non-translated mRNAs only translated after recovery from stress (Nover
et al., 1983; 1989). This suggested that the formation of HSGs protected the mRNA from the
damage caused by stress conditions, storing it until recovery to enter translation again. Later,
these similar granules containing heat shock protein HSP28 were also observed in the
cytoplasm of heat-shocked mammalian cells (Arrigo, 1990; Arrigo et al., 1988).
Subsequently, several studies have demonstrated that SGs are formed in many types of plant
and animal cells in response to a variety of stress factors including radiation (Moeller et al.,
2004), hypoxia (Gardner, 2008; van der Laan et al., 2012), oxidative stress (Emara et al.,
2012; Kedersha et al., 1999), osmotic stress (Dewey et al., 2011), nutrient stress (Tattoli et
al., 2012), some physiological conditions (Towers et al., 2011), chemical inhibitors of
translation initiation factors (Fournier et al., 2010; Mazroui et al., 2006) and during viral
infections (Lloyd, 2012). Kedersha et al. (1999) demonstrated using mammalian cells that the
RNA-binding proteins T-cell intracellular antigen 1 (TIA-1) and T-cell restricted intracellular
antigen-related protein (TIAR) colocalize with poly (A)-binding protein (PABP) in SGs
formed in response to environmental stress and suggested that the assembly of SGs depends
on the phosphorylation of eukaryotic initiation factor 2α (eIF2 which is enhanced by TIA-1
(Kedersha et al., 1999).
In mammalian cells, studies using cochlear hair cells identified that aminoglycoside
treatments using gentamycin induce TIAR translocation from the nucleus and accumulation
as punctate granules in the cytoplasm (Mangiardi et al., 2004). Moreover, the formation of
Caprin 1-containing stress granules in cochlear hair cells following aminoglycoside-induced
damage was also reported (Towers et al., 2011). Furthermore, the assembly of SGs was found
to be associated with Fragile X mental retardation (FMR) or Fragile X syndrome (FXS), and
the FMR protein (FMRP) which associated with polysomes in normal cells but in stressed
6
cells was found to colocalize with non-translated mRNA in SGs (Kim et al., 2006; Mazroui
et al., 2002). These studies have revealed that the inhibition of protein synthesis is critical for
the induction of SGs formation (Kedersha & Anderson, 2007). Chemical agents that induce
translation termination such as puromycin promote SGs formation (Kedersha et al., 2000).
Supporting this, chemical agents that arrest translation elongation and prevent polysome
disassembly, such as emetine and cycloheximide, also block SGs assembly (Kedersha et al.,
2000).
An increasing number of RNA-binding proteins have been identified as components
of stress granules using immunofluorescence studies, but the exact composition of stress
granules remains unknown and varies according to the stress, cell type or experimental
system (Anderson & Kedersha, 2008; 2009b; Buchan & Parker, 2009; Thomas et al., 2011).
For example, in yeast cells, the SGs induced under acute stress are similar in composition to
mammalian SGs. In contrast, yeast cells exposed to glucose starvation form SGs that are
distinct from mammalian SGs, lacking 40S ribosomal subunits and eIF3 (Buchan et al., 2008;
Hoyle et al., 2007). In Saccharomyces cerevisiae, NaN3-induced SGs contain the initiation
factors eIF3, eIF4A/B, eIF5B and eIF1A, whereas SGs induced by glucose deprivation do not
(Buchan et al., 2011).
1.3.1 Stress granule composition
SGs are formed within 15-20 min of stress exposure by the rapid accumulation of
transient mRNP aggregates, in the cytoplasm of cells responding to stress conditions. In
addition to non-translated poly-adenylated mRNAs, in a complex with PABP, SGs contain
translation initiation factors and a large numbers of RNA-binding proteins involved in
various functions ranging from translation, antiviral response to RNA localisation (Kedersha
7
& Anderson, 2009) (Table 1). Typically, SGs contain non-translated mRNA, 40S ribosomal
subunits (but not 60S), the eIF4F complex (eIF4E, eIF4G, eIF4A) and eIF4B, Poly(A)
binding protein (PABP), eIF3, and eIF2 (Anderson & Kedersha, 2006; Kedersha et al., 2000;
Kedersha et al., 1999; Kimball et al., 2003; Low et al., 2005; Mazroui et al., 2006). The most
characterized RNA-binding proteins are G3BP1 and TIA-I/TIAR which possessing self-
interacting domains that promote SGs assembly (Gilks et al., 2004; Tourriere et al., 2003)
and many other RNA-binding proteins regulating mRNA structure and functions. These
include proteins involved in mRNA silencing such as Argonaute proteins (Ago2 and Ago2)
(Leung et al., 2006), pumilio 2, Rap55, Staufen, TIA-1 and TIA-1-related protein (TIAR)
(Anderson & Kedersha, 2008). Proteins contributing to the antiviral response, including
APOBEC3G and APOBEC3F, are also detected in SGs supporting the role of SGs in defence
against viral infection (Gallois-Montbrun et al., 2007; Kozak et al., 2006). In addition to
translation factors, other proteins associated with mRNA translation are reported as SG
components such as Ataxin-2 (Nonhoff et al., 2007), FAST (Kedersha et al., 2005), FMRP
and FXR1 (Antar et al., 2005). Proteins involved in metabolic signalling pathways such as
G3BP1 (Tourriere et al., 2003), IP5K (Brehm et al., 2007) and TRAF2 (Kim et al., 2005),
have also been detected. Proteins involved in mRNA decay (p54/RCK/DDX6, TTP, BRF-1,
Xrn1, and Dcp1/2) (Kedersha et al., 2005; Rothe et al., 2006; Stoecklin et al., 2002;
Wilczynska et al., 2005), protein enhancing mRNA stability (HuR) (Gallouzi et al., 2000),
mRNA localisation proteins (ZBP1) (Stohr et al., 2006), and proteins important for
transcription (Rpb4 and SRC3) (Lotan et al., 2005), or splicing (MLN51) (Baguet et al.,
2007), have also been found to act as SG components. This evidence regarding the wide
nature of SG components may reflect the many possible roles for these granules in mRNA
metabolism or life cycle (Anderson et al., 2014; Buchan, 2014).
8
Table 1.1 Components of stress granules
Protein Known name Function
Argonaut proteins Ago1 Gene silencing
Ago2 siRNA , slicer
Apolipoprotein B mRNA editing enzyme,
catalytic polypeptide-like 3G
APOBEC3G Cytidine deaminase , antiviral
Cytoplasmic polyadenylation element-
binding protein 1
CPEB1 Translational silencer
Eukaryotic initiation factor 4E eIF4E Translation initiation
Fas-activated serine/threonine
phosphoprotein
FAST Signalling
Human antigen R HuR Stability, splicing
Importin 8 Imp8 Shuttling, silencing
Lin28 Lin28 Block let-7 processing
LINE 1 ORF1p
Sm-like protein1 Lsm1 mRNA splicing
hmex-3B hmex-3B Germline development
Musashi Musashi translational control that regulate
multiple stem cell populations
DEAD box protein RCK/p54 RCK/ p54 Translation /decay
RNA-associated protein 55 RAP55/Lsm14 shuttling
Roquin proteins Roquin E3 ligase, promote mRNA
degradation by recruiting the
Ccr4-Caf1-Not deadenylase
complex
Smaug protein Smg Translational silencer
T-cell intracellular antigen 1/ T-cell
restricted intracellular antigen-related
protein
TIA-1/TIAR Splicing, translational silencer
Tristetraprolin/Butyrate response factors
1
TTP/BRF-1 mRNA decay
Exoribonuclease Xrn1 mRNA decay (5' 3' exonuclease
Y box binding protein 1 YB-1 mRNA chaperone
A-kinase anchor protein 350A AKAP350A Protein scaffold
apolipoprotein B mRNA-editing enzyme
1
APOBEC1 RNA editing
Calreticulin calregulin,
CRP55, CaBP3,
Cell division cycle and apoptosis
regulator protein 1
CCAR1 Transcriptional co-activator
Caprin-1 Caprin-1 Cell cycle
Cell division cycle and apoptosis
regulator protein 1
CCAR1 Transcriptional co-activator
Caprin-1 Caprin-1 Cell cycle
Cold-inducible RNA-binding protein CIRBP Translational silencer
Death-associated protein 5 DAP5/p97/NAT1 Translation
9
Protein Known name Function
DEAD box protein A DBPA Splicing
ATP-dependent RNA helicase 1 DDX1 ds DNA breaks
ATP-dependent RNA helicase 3 DDX3 RNA helicase
Eukaryotic initiation factor 1 eIF1 Translation initiation
Eukaryotic initiation factor 2 α eIF2α Translation initiation
Eukaryotic initiation factor 3 eIF3 Translation initiation
Eukaryotic initiation factor 4A,B eIF4A, eIF4B Translation initiation
Eukaryotic initiation factor 4G eIF4G Translation initiation
Eukaryotic initiation factor 4H eIF4H Translation initiation
Ewing sarcoma protein EWS oncogene
Focal Adhesion Kinase FAK Motility, signalling
Fragile mental retardation protein / Fragile
X mental retardation syndrome-related
protein 1
FMRP/FXR1 Translation, splicing,
microRNA
Fused in Sarcoma FUS Motility
Folate-binding protein /KH-type splicing
regulatory protein
FBP/KSRP mRNA decay, miRNA
processing
Growth factor receptor-bound protein 7 GRB7 Protein scaffold
Ras GTPase-activating protein-binding
protein
G3BP Helicase, protein scaffold
Histone deacetylase 6 HDAC6 Deacetylase stress signalling
heterogeneous nuclear ribonucleoprotein A1 hnRNPA1 Splicing , multifunctional
heterogeneous nuclear ribonucleoprotein K hnRNPK mRNA processing
heterogeneous nuclear ribonucleoprotein Q hnRNPQ Splicing
Heat shock protein 27 HSP27 Heat shock
Heat shock protein 90 HSP90 Molecular chaperon
Intracellular localization of human
Ins(1,3,4,5,6)P5 2-kinase
IP5K Ins Protein scaffold
Muscleblind-like protein 1 MBNL1 Alternative splicing
Metastatic lymph node 51 MLN51 Splicing
Ploy A binding protein PABP Translation
Protein activator of the interferon-induced
protein kinase
PACT RISC, PKR activator
Plasminogen activator inhibitor 1 RNA-
binding protein
PAIRBP1
Plakophillin1/3 PKP1/3 Adhesion
plasma membrane-related Ca(2+)-ATPase 1 PMR1 Endonuclease
Pumilio 2 Pumilio 2 Development
The Receptor for Activated C Kinase 1 RACK1 Signalling, polarity
RNA binding motif protein 42 RBM42
RNA Helicase associated with AU-rich
element
RHAU helicase RNA helicase
Ribosomal s6 kinase2 RSK2 Kinase
Ribonuclease P protein subunit p20 Rpp20 RNAse P supunit
RNA polymerase II subunit B4 Rpb4 mRNA transcription
Src-associated in mitosis 68 kD SAM68 Multifunction
Stress Granule/Nucleolar Protein
SCNP Ribosome maturation
10
Protein Known name Function
The nuclear receptor coactivator 3 SRC3 Transcriptional co-activator
Staufen Staufen ds RNA binding
Survival of Motor Neuron SMN snRNP assembly
TATA-binding protein-associated factor 2N TAF15 Oncogene
Tudor domain-containing protein 3 TUDOR-3
Translin TSN Antiviral nuclease
Ubiquitin Ubiquitin Signalling protein
Z-DNA-binding protein 1 ZBP1 (zip code BP) mRNA localisation,
antiviral
The Tudor-SN protein TSN Transcription,
The energy sensor AMP-activated protein
kinase (AMPK)-2α
AMPK-α2 a central regulator of energy
homeostasis in eukaryotes,
autophagy, cell growth and
polarity to cytoskeletal
dynamics
Ataxin-2 Ataxin-2 Translation
11
1.3.2 Stress granule assembly/disassembly
Responding to stress-induced damage, eukaryotic cells activate mechanisms to save
energy and repair this damage by repressing the translation of house-keeping gene transcripts
which accumulate in SGs, and reciprocally up-regulating the translation of proteins required
to repair the cellular insults (Anderson & Kedersha, 2008). To date, the mechanism by which
stress granules assemble/disassemble is not yet fully understood, and more than 100 genes are
involved in this complex process with over 80 proteins that localise to SGs (Buchan &
Parker, 2009; Ohn et al., 2008). Many factors regulate and promote SG assembly such as
non-translating mRNAs, post-translational modifications, protein-protein interactions, and the
microtubule network (Buchan & Parker, 2009).
The first stage of SG formation is always associated with a shut-off of protein
synthesis that can be caused by different pathways (Kedersha et al., 1999; Mazroui et al.,
2006). Studies have demonstrated that one of the major mechanisms for triggering SG
formation is the phosphorylation of the alpha subunit of eIF2 on serine residue 51 (Holcik &
Sonenberg, 2005; Kedersha et al., 1999). This phosphorylation is carried out by one of four
serine/threonine kinases activated in response to different types of stress (Holcik &
Sonenberg, 2005)(Figure 1.1). The double-stranded RNA-dependent protein kinase (PKR) is
commonly activated by RNA viruses as a part of the interferon antiviral response of cell
(Maggi et al., 2000), but PKR also senses heat shock, oxidative stress and UV irradiation
(Williams et al., 2001). General control non-de-repressible-2 kinase (GCN2) senses amino
acid deprivation and the decrease of other types of nutrients, as well as UV irradiation
(Berlanga et al., 1999). PKR-like endoplasmic reticulum kinase (PERK) responds to the ER
stress caused by the accumulation of unfolded or misfolded proteins in the ER (Harding et
al., 2000; Shi et al., 1998), or hypoxia sensing (Harding et al., 2000). The heme-regulated
inhibitor kinase (HRI) is activated under conditions of intracellular iron deficiency but also
12
senses oxidative stress and heat shock (Han et al., 2001; Lu et al., 2001; McEwen et al.,
2005). Activation of all of these kinases leads to phosphorylation of eIF2, thereby
increasing the affinity (150-fold) of eIF2 for eIF2B, the eIF2 guanine nucleotide exchange
factor, blocking the guanosine diphosphate/guanosine triphosphate GDP/GTP exchange and
formation of the eIF2-GTP-tRNAMet ternary complex (TC). This prevents assembly of the
48S pre-initiation complex and results in the repression of translation initiation (Holcik &
Sonenberg, 2005; Srivastava et al., 1998).
13
Figure 1.1 Proposed model of inducting stress granules (SG) assembly by different conditions
and stimuli. The most characterised induction pathway leading to SGs assembly is the
phosphorylation of eukaryotic translation initiation factor 2α (eIF2α) by the kinases PKR, PERK, and
HRI and GCN2. This blocks the formation of eIF2-GTP-Met-tRNAiMet
ternary complex (TC) and in
turn inhibits translation initiation to trigger the assembly of SGs. In addition SGs formation can be
induced by the alteration of expression level or function of translation factors such as eIF4A, eIF4B,
eIF4H, and poly (A)-binding protein (PABP).
14
The formation of SGs can also be induced independently of the eIF2
phosphorylation pathway via cleavage or alteration, or modification of expression level and
activity, of proteins involved in translation initiation such as eIF4A, eIF4G (Anderson &
Kedersha, 2009a; Bordeleau et al., 2006; Mazroui et al., 2006), eIF4B, eIF4H, as well as
PABP (Mazroui et al., 2006). The activation or overexpression of key SG–nucleating
proteins such as G3BP1, TIA1, TIAR, TDP-43, Caprin-1, CIRP and CBEP1 also triggers the
assembly of SGs without any additional stimulus (De Leeuw et al., 2007; Gilks et al., 2004;
Kedersha et al., 2013; Tourriere et al., 2003; Wilczynska et al., 2005). However, in most
cases, it is the phosphorylation of eIF2 that leads to a rapid dissociation of polysomes even
if other routes exist (Mazroui et al., 2006).
Therefore, the first step of SG formation can occur via both eIF2 phosphorylation-
dependent and independent pathways that affect translation initiation and result in polysome
disassembly whereupon the cell begins to develop foci to store the stalled translation
initiation complexes (48S mRNPs) (Kedersha et al., 1999). Locally, RNA-binding proteins
also localise to these small foci by their affinity for the suddenly high concentration of free
mRNA in the cytoplasm (primary aggregation) (Bounedjah et al., 2014; Kedersha &
Anderson, 2009). Several factors will mediate the subsequent stages of SG assembly
mediated by proteins containing self-interaction domains that mediate protein-protein or
protein-RNA interactions (Gilks et al., 2004; Kedersha & Anderson, 2002; Tourriere et al.,
2003). Such domains are found in several RNA-binding proteins. For example, TIA-1 and
TIAR contain glutamine and asparagine-rich prion-like domains mediating SG formation via
their self-aggregating ability (Gilks et al., 2004; Kedersha & Anderson, 2002). G3BP1 also
harbours the self-interaction (polyglutamine and asparagine rich) domains, playing a critical
role in SGs assembly (Tourriere et al., 2003), and knockdown of these proteins leads to a
reduced assembly of SGs (Ghisolfi et al., 2012). PABP, which associates with most
15
transcripts, also mediates the aggregation of SGs via its C-terminal domain (Kedersha &
Anderson, 2009). Other RBPs that promote SGs assembly include FMRP/FXRI (Mazroui et
al., 2002), Staufen (Martel et al., 2010), SMN (Hua & Zhou, 2004), TTP (Stoecklin et al.,
2004), BRF1 (Kedersha et al., 2005), and CPEB (Wilczynska et al., 2005). Proteins lacking
RNA-binding features can also contribute to this process via protein-protein interaction with
certain mRNA binding proteins in SGs, so-called piggyback proteins (Kedersha & Anderson,
2009). For example, SRC3, FAST, PMRI are localized to SGs via interaction with TIA-1
(Yang et al., 2006b; Yu et al., 2007). In addition, TRAF2 and plakophillin 3 that bind to
eIF4G and G3BP1, respectively, are recruited to SGs in a piggyback manner (Hofmann et al.,
2006; Kim et al., 2005). The recruitment of specific SGs proteins components during this
stage is largely associated with an excess of free mRNA (non-polysomal mRNA) and the
nucleic acid/proteins imbalance may also regulate the SGs assembly (Bounedjah et al., 2014).
Protein modifications are also involved in the assembly of SGs by enhancing the
recruitment of several proteins to SGs (Ohn & Anderson, 2010). Phosphorylation and
dephosphorylation of several proteins play a critical role in SGs assembly, for example the
phosphorylation of eIF2 plays a key role in initiation of SG formation (Kedersha et al.,
1999). The overexpression and phosphorylation of G3BP also regulate SG
assembly/disassembly, while the overexpression of G3BP1 induces SGs assembly; the
phosphorylation of this protein at serine residue 149 inhibits SGs formation (Tourriere et al.,
2003). Phosphorylation of other proteins has also been reported to regulate SGs
assembly/disassembly such as phosphorylation of growth factor receptor-bound protein 7
(Grb7) by focal adhesion kinase (FAK) in heat shock–stressed cells (Tsai et al., 2008).
Phosphorylation of tristetraprolin (TTP) regulates the interaction between SGs and PBs
(Stoecklin et al., 2004), and phosphorylation of eIF4E-transporter (4E-T) enhances stress-
dependent PBs assembly (Cargnello et al., 2012). Protein modification such as the
16
hypusination of eIF5A via a polyamine biosynthetic pathway inhibits SGs assembly (Li et al.,
2010). Moreover, studies have reported that the methylation of the RGC domain of FMRP
and cold inducible RNA-binding protein (CIRP) are required for SGs assembly (De Leeuw et
al., 2007; Dolzhanskaya et al., 2006). Furthermore, Ohn et al., (2008) observed that the O-
GlcNAc modification of ribosomal proteins correlates with SGs formation (Ohn et al., 2008).
Finally, the cytoskeleton and associated motor proteins also participate in
assembly/disassembly of SGs by transporting stalled mRNPs to sites of RNA granule
assembly (Bartoli et al., 2011; Loschi et al., 2009). The depolymerisation of microtubules
using nocodazole or vinblastine prevented the assembly of SGs in a mammalian CV-1 cell
line that was treated with arsenite, while microtubule-stabilizing drugs such as paclitaxel
promote SG formation (Ivanov et al., 2003). It was further proposed that the disruption of
microtubules affects the number and size of SGs rather than inhibiting their formation
(Fujimura et al., 2008; Kolobova et al., 2009; Loschi et al., 2009). In contrast, Kwon et al.,
(2007) reported that the stabilization of microtubules was not associated with SGs assembly
(Kwon et al., 2007). In addition to microtubules, microtubule motor proteins such as dynein
and kinesin localize to SGs and also contribute in SGs assembly/disassembly. Studies have
reported that dynein facilitates SG assembly and the absence of this protein from cells treated
with arsenite has a negative impact on the size and number of SGs (Kwon et al., 2007; Loschi
et al., 2009), while kinesin plays a role in disassembly of SGs after stress recovery (Loschi et
al., 2009). According to these findings, the role of microtubules and associated motor
proteins is to facilitate coalescence of small aggregates to form large aggregates (Kolobova et
al., 2009; Tsai et al., 2009). Furthermore, pre-existing PBs may be required for the assembly
of SGs (Buchan et al., 2008). Depending on the SGs component, the mRNAs that localise to
SGs can either exit SGs and re-enter translation, detach from the eIF3/40S complex and
17
localise to PBs for degradation via their binding to specific proteins such as TTP, or be stored
in SGs for a prolonged time (Kedersha & Anderson, 2009).
The SGs disassembly process begins when the cells recover from the stress naturally
or artificially (using drugs stabilising polysomes) and translation returns to normal (Anderson
& Kedersha, 2008; Nadezhdina et al., 2010). Other factors such as turnover of their protein
component, chaperone proteins, and autophagy are possibly involved in SG disassembly
(Buchan, 2014).
1.3.3 Functions and importance of stress granules
In addition to their role in maintaining cellular homeostasis during noxious conditions
via retention and protection of the specific mRNAs stored in a repressed state enabling the
cell to repair the cellular damage until stress recovery (Hofmann et al., 2012; Kedersha et al.,
2005; Tsai & Lloyd, 2014), SGs might participate in a variety of functions mediated by SG-
associated proteins, which include translation, decay, splicing, localization, stability,
signalling, and response to viral infections (Buchan, 2014; Kedersha & Anderson, 2009).
SG proteins TTP and BRF1/2 that bind to 3′ untranslated regions of mRNA promote
the interaction between SGs and PBs to facilitate mRNA decay within PBs (Anderson &
Kedersha, 2008; Brengues et al., 2005; David Gerecht et al., 2010; Kedersha et al., 2005;
Yang et al., 2006a). Zipcode-binding protein 1 (ZBP1) controls mRNA turnover, translation
and localisation in non-stressed cells (Yisraeli, 2005). During cellular stress, the interaction
of this protein with specific mRNAs within SGs is required for the stabilization of associated
mRNAs, regulating the mRNA fate during stress conditions (Stohr et al., 2006). Moreover,
knockdown of ZBP1 leads to destabilisation of specific mRNAs (Stohr et al., 2006). Several
18
SG proteins, including DBPA, FASTK, FMRP/FXR1, hnRNPA1, HuR, SAM68, TIA-1 and
TIAR shuttle between the nucleus and cytoplasm, and this shuttling might play a role in
alternative splicing and microRNA processing (Kedersha & Anderson, 2009). The temporary
sequestration of signalling proteins such as G3BP1, FXR1/FMRP, TIA-1/TIAR, TRAF2,
RACK1 and FAST into SGs may affect multiple signalling pathways as well as RNA
metabolism, thus regulating the survival of the stressed cell (Anderson & Kedersha, 2008;
Kedersha et al., 2013; Kim et al., 2005). Stress granules might also function as a regulator of
cell death (Mokas et al., 2009). The sequestration of pre-apoptotic proteins such as RCCK1,
ROCK1, and TRAF2, as well as mRNA encoding pro-apoptotic proteins inhibits the
apoptotic response (Arimoto et al., 2008; Kim et al., 2005; Moeller et al., 2004). In addition,
the disruption of RNA granule function is associated with several diseases including
neurodegenerative diseases and dementias, immunological and viral diseases (Bloch et al.,
2013; Buchan, 2014; Lloyd, 2013). These diseases usually correlate with mutation of proteins
found as components of SGs such as FMRP, fused in sarcoma (FUS), transactive response
(TAR) DNA binding protein-43 (TDP-43) and ataxin-2 (Anderson & Kedersha, 2008;
Shelkovnikova et al., 2013). For example, Fragile X syndrome (FXS) results from mutations
within the X-linked FMR1 gene which lead to neuronal cell death and may be correlated with
the inhibition of SG assembly (Didiot et al., 2009; Kim et al., 2006). It was proposed that
impairing SG assembly prevents the reprogramming of SG FMRP protein translation (that
protect the cells from sudden environmental stress) leading to FXS (Didiot et al., 2009; Kim
et al., 2006). Two SG proteins, TDP-43 and FUS/TLS, which harbour prion-like domains,
play critical roles in amyotrophic lateral sclerosis (ALS), a neurodegenerative disease
primarily affecting motor neurons (Cleveland & Rothstein, 2001). Mutations in TDP-43
protein increase the tendency of this protein to form SGs in response to stress as well as its
binding to other proteins within SGs, such as TIA-1 or FUS (Liu-Yesucevitz et al., 2010).
19
The sequestration of these proteins in SGs reduces their level in the nucleus and
neurodegeneration could arise from either low levels of nuclear FUS and TDP-43 or
increased formation of SGs (Anderson et al., 2014). SGs have also been proposed to be
involved in the pathogenesis of other diseases such as spinal motor atrophy (Hua & Zhou,
2004), ischemia–reperfusion (Kayali et al., 2005), Huntington’s chorea and Alzheimer’s
disease (Vanderweyde et al., 2013). SGs and PBs are also involved in the mechanisms used
by cancer cells to re-program gene expression and enhance cell survival by modulating
cellular signalling pathways, metabolic machinery, and stress response programs (Anderson
et al., 2014). Several studies have reported the involvement of RNA granules in cancer
development as well as demonstrating an interplay with the response to chemotherapy and
specific drugs (Anderson et al., 2014). For instance, an association between
chemotherapeutic treatments using bortezomib and increased SGs formation was found to be
necessary for cancer cell survival. Using different types of cancer cell line, including colon
cancer (Caco), cervical cancer (HeLa) and lung cancer (Calu-I) cell lines, formation of SGs
was found to be associated with the ability to resist bortezomib-mediated apoptosis, (Fournier
et al., 2007). The antimetabolite 5-fluorouracil (5-FU) used for treatment of different types of
cancer such as breast, colorectal, head and neck cancer (Longley et al., 2003), also induces
eIF2α phosphorylation-dependent-SGs through activation of PKR leading to cell survival via
the sequestration of RACK1, a protein mediating cell survival and apoptosis (Kaehler et al.,
2014; Longley et al., 2003). Other types of drugs such as Pateamine A (Pat A) (Bordeleau et
al., 2005), hippuristanol (Bordeleau et al., 2006), and silvestrol (Bordeleau et al., 2008),
affecting eIF4A activity and leading to translation initiation inhibition, induce SG assembly
and are anti-tumour agents in many types of cancer (Anderson et al., 2014).
In recent years, there has been growing body of evidence to indicate that SGs and PBs
form part of the cellular response to stress triggered by viral infection (Beckham & Parker,
20
2008; Lloyd, 2013) (see section 1.6). As discussed above, the importance of RNA granules is
not just as a means of storage or degradation of stalled mRNAs, but also because of their
influence on several aspects of cell metabolism and gene regulation during abnormal or
unsuitable conditions (Anderson et al., 2014).
1.4 Processing bodies (P-bodies or PBs)
PBs are the second major type of cytoplasmic RNA granules associated with stressed
or unstressed somatic cells of all eukaryotes such as vertebrates, invertebrates, and plants as
well as in yeast and trypanosomes; and their function is conserved from yeast to human (Jain
& Parker, 2013b; Kedersha & Anderson, 2009; Kulkarni et al., 2010; Parker & Sheth, 2007).
The presence of PBs is constitutive in the cytoplasm of unstressed cells, but their number and
size can increase several fold in response to stress (Anderson et al., 2014; Jain & Parker,
2013b; Kulkarni et al., 2010; Parker & Sheth, 2007; Teixeira et al., 2005). The presence
within PBs of enzymes required for mRNA degradation and surveillance, translational
repression, and RNA silencing might reflect their association with translation repression
and/or mRNA decapping and degradation (Baguet et al., 2007; Brengues et al., 2005; Decker
& Parker, 2012; Decker et al., 2007; Parker & Sheth, 2007). PBs were first observed by
Bashkirov in 1997 as a punctate granular signal of the mammalian Xrn1 exoribonuclease, the
main cytoplasmic 5′→3′ exoribonuclease in eukaryotic cells, in the cytoplasm of a mouse
fibroblast cell line and thus named “Xrn1 foci” (Bashkirov et al., 1997). Then, Eystathioy in
2002 observed a novel protein, GW182, localised to these foci in the cytoplasm of patients
suffering from motor and sensory neuropathy, naming this structures as GW182 bodies
(Eystathioy et al., 2003). Later, other proteins involved in decay mechanism were found to
localise to PBs, including the decapping enzymes Dcp1/Dcp2 (Ingelfinger et al., 2002), the
21
decapping activators Hedls (Yu et al., 2005), Edc3 (Fenger-Gron et al., 2005), Pat1 (Scheller
et al., 2007), Lsm1-7 (Ingelfinger et al., 2002), and RCK (p54, DDX6) (Wilczynska et al.,
2005). Due to the presence of RNA decay machinery components, these cytoplasmic foci are
also called ‘decapping-bodies’ or ‘mRNA decay bodies’, but the most widely accepted name
is P-bodies or PBs (Sheth & Parker, 2003). Moreover, PBs are enriched with other types of
proteins such as RNA degradation proteins, the deadenylase complex CCR4/CAF1/NOT
(Zheng et al., 2008), its enhancer TOB2 (Ezzeddine et al., 2007), nonsense-mediated mRNA
decay proteins UPF1, UPF2, UPF3, SMG5, SMG6 and SMG7, ARE-mediated decay factors
TTP, BRF1 and BRF2 (Franks & Lykke-Andersen, 2007; Kedersha et al., 2005; Stoecklin &
Anderson, 2007), and the RNAi machinery component Argonaut (Sen et al., 2005). In
addition, PBs contain protein involved in translation regulation such as eIF4E (Andrei et al.,
2005), eIF4E-T (Ferraiuolo et al., 2005), FASTK (Kedersha et al., 2005), CPEB (Wilczynska
et al., 2005), and PCBP2 (Fujimura et al., 2008). Table 1.2 shows several other proteins that
can be detected as components of PBs. Moreover, PBs contain translationally repressed
mRNA in response to a variety of stresses such as oxidative stress, UV stress, osmotic shock
and nutritional deprivation (Buchan et al., 2008; Buchan et al., 2011; Izawa et al., 2007;
Teixeira et al., 2005; Yoon et al., 2010).
22
Table 1.2 Components of P-bodies
Protein Known name Function
C-C chemokine receptor type 4 CCR4 Transcription, ribosome
biogenesis
cytoplasmic polyadenylation element binding
protein
CPEB Translation regulator
Decapping enzymes Dcp1and Dcp2 mRNA decay
Enhancer of decapping protein 1,2 Edc1,2 mRNA decay
Enhancer of decapping protein 3 Edc3 mRNA decay
Eukaryotic initiation factor 4E eIF4E Translation initiation
Eukaryotic initiation factor transporter 4E-T eIF4E-T mRNA decay
GW182/Trinucleotide repeat-containing gene
6A protein
GW182/TNRC6B siRNA
Guanylate binding protein 2, interferon-
inducible
Gbp2 mRNA export
Eukaryotic initiation factor 4G eIF4G Translation initiation
Hedls/Ge-1 Hedls/Ge-1 Enhancing mRNA
decapping
Heterogeneous nuclear ribonucleoprotein A3 hnRNPA3 Splicing, multifunctional
Importin-8 Importin-8
Putative helicase MOV-10 MOV10/Armitage RNA-directed transcription
NFX7 NFX7 mRNA export
Protein interacting with APP tail-1
Pat1/Pat1L Decapping
CCR4–CAF1–NOT complex CCR4–CAF1–NOT
complex
Deadenylation
Regulator of nonsense transcripts 1,2,3 UPF1,2,3 Nonsense-mediated decay
(NMD)
Eukaryotic release factor 1 and 3 eRF1 and eRF3 Translation termination
exonuclease Xrn1 5′ to 3′ exonuclease
Rap55 Rap55/Scd6 Translation repressor
Fas activated serine/ threonine
phosphoprotein FAST
Poly C- binding protein 2 PCBP2 Translation regulator
23
1.4.1 P-bodies assembly/disassembly
Assembly of PBs is initiated when translation is repressed in response to a cellular cue
by the interaction of non-translated mRNA with various proteins forming mRNPs, followed
by the aggregation of these mRNPs via self-interaction domains leading to the formation of
PBs visible under microscopy (Decker et al., 2007; Eulalio et al., 2007a). Generally, this
aggregation is reversible, and mRNPs can re-enter translation, or in some cases mRNPs
undergo further rearrangement via association with specific proteins that facilitate the decay
or degradation of mRNA (Bhattacharyya et al., 2006; Brengues et al., 2005; Eulalio et al.,
2007a). Whether the formation of PBs is a de novo process or the mRNA and associated
proteins are localised to pre-existing structures remains an unresolved issue (Jakymiw et al.,
2007).
The mechanism of PB assembly is not entirely understood and varies depending on
the cell system (Decker et al., 2007; Franks & Lykke-Andersen, 2007; 2008). More than 40
genes have been proposed to be involved in the assembly process (Ohn et al., 2008). The
presence of mRNA is required for formation and integrity of PBs (Andrei et al., 2005;
Eystathioy et al., 2002; Teixeira et al., 2005). Treatments with cycloheximide, which traps
the mRNAs in polysomes, or with RNase both lead to the disappearance of visible PBs in
mammalian cells (Cougot et al., 2004; Eulalio et al., 2007a; Franks & Lykke-Andersen,
2008; Teixeira & Parker, 2007). In addition to repressed mRNA, specific PB proteins are
critical for the formation of a small core during PB assembly, and subsequently many other
proteins contribute to this process (Decker et al., 2007; Eulalio et al., 2007a; Kulkarni et al.,
2010; Liu et al., 2005; Yu et al., 2005).
24
In yeast, three proteins are essential for assembling PBs: Edc3 (Decker et al., 2007;
Kulkarni et al., 2010), Pat1 (Pilkington & Parker, 2008), and Lsm4 (Ingelfinger et al., 2002).
All of these proteins possess multi-domains that mediate the assembly process via self-
interaction properties and interacting with other proteins. Three separate domains of Edc3
mediate the self-interaction and its association with other proteins including Dcp1 and the
RNA helicase Dhh1/RCK leading to an enhanced PB assembly (Decker et al., 2007; Kulkarni
et al., 2010). The Pat1 C-terminal domain promotes PB assembly via its interaction with
DCP1, while the middle domain of Pat1 interacts with Lms1 (Pilkington & Parker, 2008). In
addition, the Q/N-rich prion-like domain of Lsm4p is required for P-body assembly (Decker
et al., 2007). Decker et al, 2007 suggested that two sub-complexes of a conserved core of
proteins associate directly with mRNA playing an important role in the assembly of PBs, the
first complex consists of Dcp1p, Dcp2p, Dhh1p, and Edc3p and the second consists of Pat1p,
Lsm1-7p, and Xrn1p. The interaction between these two complexes via Edc3p YjeF-N
domain and/or the Lsm4p Q/N-rich prion-like domain is proposed to mediate PB formation
(Decker et al., 2007). Moreover, the YjeF-N domain is conserved amongst higher eukaryotes
promoting PB assembly (Ling et al., 2008). In metazoan, Lsm4 Q/N-rich prionlike domain is
not conserved and instead is replaced by Arg-Gly-Gly (RGG) involved in the SG assembly
process (Tourriere et al., 2003). Other proteins containing a Q/N region have been identified
as metazoan PB components including GW182 and Hedls/Ge-1 protein (Decker et al., 2007;
Eulalio et al., 2007b; Liu et al., 2005; Yu et al., 2005). In addition to GW182, other proteins
including RAP55 (also known as Lsm14) and Ge-1 (also known as Hedls or RCD-8) are
required for PB assembly in human cells, and depletion of these proteins leads to the
disassembly of PBs (Eulalio et al., 2007b; Yang et al., 2006b; Yu et al., 2005). In
mammalian cells, PB formation is greatly reduced by the depletion of GW182 and Ge-1,
which have no orthologous in yeast (Eulalio et al., 2007a), as well as Lsm1, RCK/p54 (Dhh1)
25
(Chu & Rana, 2006; Coller & Parker, 2005), and eIF4E-T (Andrei et al., 2005; Ferraiuolo et
al., 2005). Moreover, the disassembly of PBs is associated with the inhibition of miRNA
pathway, and the depletion of proteins involved in miRNA process, such as Drosha and its
binding partner DGCR8, lead to the dissolution of PBs (Pauley et al., 2006). PBs are highly
mobile structures that move along microtubules and the link between the intracellular
microtubule network and formation of PBs has been reported (Aizer et al., 2008; Aizer &
Shav-Tal, 2008; Baguet et al., 2007; Sweet et al., 2007). Aizer et al., 2008 demonstrated that
disrupting microtubules using the drug benomyl caused aggregation of PB components in
yeast (Sweet et al., 2007), which suggested that the enhancement of PB formation may occur
by preventing PB components from being trafficked away by microtubules. The same
relationship between cytoskeleton and assembly of PBs is also reported in mammalian cells
(Aizer et al., 2008; Aizer & Shav-Tal, 2008). These studies suggested that the formation of
PBs is controlled under normal conditions and that microtubules are involved in this process
to restrict the size or number of PBs. Furthermore, the assembly of RNA granules is
controlled by different signalling pathways. For example, the PKR pathway plays an
important role in the regulation of PB assembly/disassembly in yeast by phosphorylating the
Pat1 protein (Shah et al., 2014).
1.4.2 P-bodies function
The localisation of proteins associated with processes such as mRNA decay,
surveillance, silencing and translation repression in PBs has encouraged many researchers to
investigate any possible role for these cytoplasmic foci in these mechanisms. However, the
role of PBs varies depending on the particular cell system and stress condition that induces
these cytoplasmic granules (Shah et al., 2013).
26
1.4.3 Role of P-bodies in mRNA degradation
Eukaryotic cells employ two pathways for degrading mRNAs, both of which start
with shortening of the 3′ poly (A) tail (deadenylation) mediated by the Pan2-Pan3 and Ccr4-
Not complexes. Following deadenylation, mRNA can either undergo degradation from 5′-3′
by Xrn1, the 5′-3′ mRNA decay pathway, or degradation by the exosome from 3′-5′ after
removal of the 5′ cap structure by decapping enzymes, the 3′-5′ decay pathway (Balagopal &
Parker, 2009; Chen & Shyu, 2011; Coller & Parker, 2004; Wahle & Winkler, 2013)(Figure
1.2). To date, the actual role of PBs in mRNA decay is not yet clear, but the presence of 5′-3′
mRNA decay pathway enzymes, such as Dhh1, Xrn1, Dcp1/2, Pat1, Edc3, Lsm1-7 and Ccr4-
Not, strongly supports that these structures are potential sites for mRNA degradation (Jain &
Parker, 2013a). Several observations provide evidence to support this idea. For instance, the
appearance of PBs is associated with the presence of mRNA, and blocking transcription using
Actinomycin D or blocking mRNA decay by preventing deadenylation both affect the
assembly of PBs (Andrei et al., 2005; Cougot et al., 2004; Teixeira et al., 2005). Moreover,
the recognition of mRNA decay intermediates in PBs also supports the role of PBs in mRNA
decay (Sheth & Parker, 2003). The exosome and its Ski co-factors have not been identified as
PB components and therefore PBs are not associated with 3′-5′ degradation of mRNA
(Brengues et al., 2005; She et al., 2004). Furthermore, the PB components Pat1 and Lsm1-7
play a role in limiting exosome-mediated 3′-5′ mRNA degradation (He & Parker, 2001).
27
Figure 1.2 Eukaryotic mRNA decay pathways. The two general decay pathways in
eukaryotes include 3′ to 5′ and 5′ to 3′ degradation, both of which start with the
deadenylation of the 3′ poly (A) tail of the mRNA by the deadenylase
Ccr4p/Pop2p/Not complex. The deadenylation is followed by either 3′ to 5′
degradation by the exosome or by decapping by the 5’ decapping, mediated by
Dcp1/Dcp2 and 5′ to 3′ degradation by the exonuclease Xrn1. This second pathway is
thought to occur in P-bodies.
28
1.4.4 Role of P-bodies in mRNA surveillance
Nonsense-mediated decay (NMD) is one of the quality control mechanisms employed
by eukaryotic cells to terminate the translation of aberrant mRNA (Baker & Parker, 2004).
The surveillance complex involved in NMD includes UPF1-3 proteins and NMD effectors
SMG1/ 5/6/7 that induce termination of premature translation (Behm-Ansmant & Izaurralde,
2006). This complex is involved in the recruitment of functional proteins that promote
mRNA decay such as Xrn1 and decapping enzymes in yeast (Cao & Parker, 2003). The link
between mRNA surveillance and PBs is not fully clear, but data suggest that SMG7 localises
to PBs and induces the aggregation of SMG5 and UPF1 in these foci (Fukuhara et al., 2005;
Sheth & Parker, 2006). UPF1 also promotes the accumulation of UPF2 and UPF3 in PBs of
yeast cells lacking Dcp1 (Eulalio et al., 2007a). The accumulation of NMD proteins in PBs in
yeast may reflect the possible role for these RNA foci in mRNA surveillance (Eulalio et al.,
2007a). By contrast, Stalder and Mühlemann, 2009 have reported that the NMD process in
mammalian cells does not require PBs (Stalder & Muhlemann, 2009). PBs have also been
linked to [AU-rich element (ARE)-mediated mRNA decay] AMD, a decay process involving
ARE-containing mRNAs. TTP/BRF proteins may also mediate this degradation process (Lai
et al., 2000; Stoecklin et al., 2002). Depletion or knockdown of general decay pathway
proteins, Xrn1, Lsm1, Dcp1 or 4E-T all lead to inhibition of AMD and accumulation of ARE-
containing mRNAs and TTP/BRF proteins in PBs (Ferraiuolo et al., 2005; Franks & Lykke-
Andersen, 2007; Stoecklin et al., 2006).
1.4.5 Role of P-bodies in gene silencing
Small RNAs (~22 nt), including small interfering RNAs (siRNAs) and microRNAs
(miRNAs), involved in gene silencing, employ different effector mechanisms to silence their
29
target genes (Liu et al., 2005). This process is involved in post-transcriptional gene
expression regulation in higher eukaryotes (van Wolfswinkel & Ketting, 2010). The role of
PBs in RNA-mediated gene silencing is still a controversial issue (Leung & Sharp, 2013).
The localisation of specific RNA-induced silencing complexes (RISC) proteins such as
Argonaute proteins to PBs supports a role for PBs in RNA silencing (Liu et al., 2005).
Studies have demonstrated that Argonaute proteins also interact with other PB components
such as GW182, Dcp1, Dcp2 and RCK/p54, suggesting the presence of these proteins as a
complex in PBs (Eulalio et al., 2007a; Jakymiw et al., 2005; Liu et al., 2005). In human cells,
the depletion of GW182 or of the Dcp1-Dcp2 complex both impair miRNA function (Liu et
al., 2005). However, the depletion of other PB components such as Lsm1 or RCK/p54,
leading to loss of PBs has no effect on siRNA function (Chu & Rana, 2006; Jagannath &
Wood, 2009). Disruption of PB formation by trapping the mRNA in polysomes also did not
correlate with siRNA function (Chu & Rana, 2006; Franks & Lykke-Andersen, 2007).
1.4.6 Other functions of P-bodies
PBs are found to be important for the long-term survival of the cell and are present in
stationary phase in a process mediated by the PKA signalling pathway (Ramachandran et al.,
2011; Shah et al., 2013). PBs have also been proposed to be involved in the immunity
disorder (Bloch et al., 2013). A link between PB formation and translational repression or
regulation (Coller & Parker, 2005), and mRNA storage (Brengues et al., 2005; Bhattacharyya
et al., 2006) during viral infection has also been extensively reported (Tsai & Lloyd, 2014)
(see later section).
30
1.5 Interactions between stress granules and P-bodies
Despite differences between SGs and PBs in their composition (SGs contains 40S
ribosomal subunits and associated translation initiation factors such as eIF3 but these are not
present in PBs), size (PBs 100-300 nm and SG 0.2-5 μm), and mechanism of assembly and
function, a dynamic link between these cytoplasmic granules has been reported (Anderson &
Kedersha, 2006; 2009a; Kedersha et al., 2005; Wilczynska et al., 2005). SGs and PBs can
share certain proteins (such as eIF4E, RCK, TIA-1, TIAR, TTP, Xrn1 and Argonaute
proteins), making a clear distinction between them difficult, and they also contain specific
translationally repressed mRNA (Anderson & Kedersha, 2008; Buchan et al., 2008; Kedersha
et al., 2005; Mokas et al., 2009) (Figure 1.3). Although, direct evidence of mRNA transit
between these granules is still lacking, some studies have reported that the overexpression of
certain proteins such as TTP or BRF1 (Kedersha et al., 2005), and CPEB1 (Wilczynska et al.,
2005), mediated mRNA transit between these two cytoplasmic granules within the same cell.
This may reflect the presence of the same reporter mRNA in these foci, and these structures
would house mRNPs representing different stages of the mRNA life cycle rather than
different types of transcripts (Anderson & Kedersha, 2009a). Both granules form in the
cytoplasm of all eukaryotic somatic cells in response to several types of stress and their
formation is strongly associated with disassembly of polysomes (Hofmann et al., 2012;
Kedersha et al., 2005). Buchan et al., 2008 demonstrated that the SGs appear next to PBs in
S. cerevisiae (Buchan et al., 2008). In mammalian cells, the docking of these granules
together was also detected using live cell imaging experiments (Kedersha et al., 2005).
Furthermore, the live cell imaging also revealed an association and fusion events between
PBs and SGs (Kedersha et al., 2005; Wilczynska et al., 2005). According to these findings,
these cytoplasmic granules can transiently interact together (Anderson & Kedersha, 2009a).
31
Figure 1.3 Components of P-bodies and stress granules. The proteins markers found in
stress granules only (red), P-bodies only (yellow) and in both are displayed.
32
Because of the reverse correlation between disassembly of polysomes and assembly
of SGs or PBs and the dynamic link between these granules, as well as the ability of mRNA
to re-enter translation following relief from stress (Anderson & Kedersha, 2006;
Bhattacharyya et al., 2006; Brengues et al., 2005), several studies have suggested that the
eukaryotic mRNAs are in dynamic equilibrium between a translated pool (polysomes) and
non-translated pool either stored in SGs or degraded in PBs (Anderson & Kedersha, 2009a;
b; Balagopal & Parker, 2009; Brengues et al., 2005; Decker & Parker, 2012; Layana et al.,
2012; Lloyd, 2012) (Figure 1.4). Thus the mRNA life cycle between these subcellular
structures can affect the rate of mRNA translation and degradation (Beckham & Parker,
2008). Recently, Simpson et al., 2014 investigated the re-distribution of specific mRNA to
PBs after stress in live yeast cells using the mTAG system and reported that the movement of
specific endogenous mRNAs is biphasic, with some mRNAs present in RNA granules early
after stress, while other mRNAs localise much later in the stress period and these events were
associated with the recruitment of specific PB proteins such as eIF4E and Bfr1p (Simpson et
al., 2014). Distinguishing which mRNA species are present in SGs or PBs remains a matter
of debate (Thomas et al., 2011; Tsai & Lloyd, 2014). Available data reported the exclusion of
certain stress-active transcripts such as heat shock protein mRNA from SGs during heat stress
(Anderson & Kedersha, 2002; Nover et al., 1989). Other studies have confirmed the presence
of certain mRNA transcripts in SGs (Mazroui et al., 2007; Stohr et al., 2006). Many factors
may be involved in determining which mRNA transcripts will be targeted to RNA granules,
for example: the presence of proteins that harbour U-rich RNA such as TIA-1 (Buchan &
Parker, 2009; Cassola et al., 2007), which bind specific mRNA targeting them to SGs. More
recently, Lui et al., 2014 using live imaging of yeast cells, showed the localisation of specific
glycolytic mRNAs to cytoplasmic granules in unstressed growing cells. Following stress,
these granules coalesce and serve as a platform for PB assembly through the recruitment of
33
mRNA decay components. Interestingly, these mRNAs are translated within these
cytoplasmic granules under non-stress conditions and not associated with the mRNA decay
machinery. This suggests that the storage or decay of some mRNAs is driven by the pre-
existence of mRNP granules containing translationally active mRNAs (Lui et al., 2014).
Unlike PBs and SGs which harbour repressed mRNAs, the mRNA granules of unstressed
cells were unaffected by cycloheximide treatment which traps mRNAs in polysomes,
indicating that the ribosome may continue translation elongation in these granules (Lui et al.,
2014).
34
Figure 1.4 Proposed model for the role of cytoplasmic RNA granules in mRNA fate.
The mRNA is in dynamic equilibrium between a translated state (polysome) and un-
translated state (SGs and PBs). Under normal condition, the mRNA associates with
polysomes and is translated into polypeptides. Following stress, mRNAs can be stored in
cytoplasmic granules (SGs or PBs) to stall their translation. mRNAs can exit the SGs or
PBs to re-enter translation upon stress recovery or be further targeted to PBs for
degradation.
35
1.6 Stress granules, P-bodies, and viral life cycles
Viruses depend on the host cell for their replication cycle as they lack the
macromolecular machinery and small molecule precursors required to complete their
replication cycle (Birch et al., 2012). Therefore a variety of mechanisms are employed by the
infected host cells to cope with and restrict viral infection (Montero & Trujillo-Alonso, 2011;
Pattnaik & Dinh, 2013; Reineke & Lloyd, 2013; Tsai & Lloyd, 2014). The formation of SGs
and PBs in response to stress generated by viral infection was recently described as a novel
cellular mechanism to restrict viral infection (Beckham & Parker, 2008), silencing translation
thereby giving the cells more time to decide their fate via activation of other antiviral
signalling pathways as well as conserving their energy while accumulating proteins that are
required for adaptation (Pichon et al., 2012; Simpson & Ashe, 2012; Tsai & Lloyd, 2014). In
parallel, viruses have evolved different strategies to counteract, tolerate, or even take
advantage of these antiviral responses via the modulation of SG and PB formation during
infection (Beckham & Parker, 2008; Montero & Trujillo-Alonso, 2011; Schutz & Sarnow,
2007). Viruses can affect RNA granule formation either by disrupting the assembly of RNA
granules or utilizing RNA granule proteins for viral propagation (Reineke & Lloyd, 2013).
The understanding of the interplay between PBs, SGs and viral life cycle is still recent and
developing (Beckham & Parker, 2008; Lloyd, 2013; Reineke & Lloyd, 2013).
A large body of work supports this relationship through the identification of critical
proteins within PBs or SGs that promote or restrict viral infection, or by the demonstration
that some viral RNAs and proteins, as well as host antiviral defence proteins, accumulate in
PBs and/ or SGs (Beckham & Parker, 2008; Jain & Parker, 2013a; Kedersha & Anderson,
2007; 2009; Lloyd, 2013; Reineke & Lloyd, 2013; White & Lloyd, 2012). The sequestration
of mRNA in a translationally silenced state in both SGs and PBs affects viral mRNA fate;
therefore viruses have evolved a variety of mechanisms to avoid this fate (Lloyd, 2012). The
36
broad range of virus-RNA granule interactions is largely dependent on the viral system and
will be further discussed within the next sections (Pattnaik & Dinh, 2013; Tsai & Lloyd,
2014).
1.6.1 Viral interaction with stress granules
Viruses interact with SGs via a variety of strategies which may generally be of two
types: either a virus-mediated blockade of SG formation or viral replication that occurs
despite the presence of SGs because the virus is able to tolerate or exploit the SG response
(Montero & Trujillo-Alonso, 2011; White & Lloyd, 2012) (Figure 1.5).
The inhibition of SGs formation by viruses can occur throughout infection or in the
mid-phase of infection depending on the virus. In both cases, the induction of SGs usually
happens early in infection following the activation of eIF2 kinases by virus recognition
(Montero & Trujillo-Alonso, 2011). Later, the inhibition of SGs formation occurs either by
cleavage/modification of SG components or by sequestering key SG-nucleating components
that drive SG assembly (Lloyd, 2012; Tsai & Lloyd, 2014). Additionally, the manipulations
of PKR or in some cases PERK also contribute to the inhibition of stress granules formation
(Lloyd, 2013). Interestingly, some viruses benefit from the SG-mediated antiviral response by
recruiting specific SG components to their replication complex to enhance their replication
cycle (Emara & Brinton, 2007; Li et al., 2002; Panas et al., 2012; Yi et al., 2011). Moreover,
a complex oscillation between SG assembly and disassembly has also been reported during
certain viral infection (Ruggieri et al., 2012).
37
Figure 1.5 Mechanisms of stress granules disruption by viruses. Viruses have evolved
several mechanisms to disrupt or prevent SGs assembly. (A) HCV, flavivirus, JUNV,
alphavirus and VSV can all induce sequestration of different SG components to their
replication complex (RC). (B) The picornavirus proteases can cleave nucleating-SG proteins
leading to inhibit SG assembly. (C) Viral proteins can interact with SGs components or SG-
inducing pathways to prevent the formation of SGs. (D) The re-localisation of SG proteins
from cytoplasmic foci to nucleus can also affect the formation of SGs as in the case of IAV
and rotavirus.
38
1.6.2 Inhibition of Stress granules by viral proteinase
Three picornaviruses including poliovirus (PV), coxsackievirus B3 (CVB3) and
encephalomyocarditis virus (EMCV) have been found to induce stress granule formation
early in infection (2-3 h.p.i), independently of eIF2 phosphorylation (Fung et al., 2013; Ng et
al., 2013; White et al., 2007). Later, the disassembly of these granules during the middle
phase of the viral replicative cycle (3-8 h.p.i) occurs via the cleavage of G3BP by the viral 3C
proteinase at the residue glutamine 326, thereby preventing the localisation of mRNA and
associated factors such as the initiation factors eIF4G and PABP and 40S ribosomal subunits
to cytoplasmic foci (White et al., 2007). In contrast, the aggregation of TIA-1 still occurs at
late times post-infection in PV-infected cells forming unconventional SGs that lack G3BP,
eIF4G and PABP (White et al., 2007; White & Lloyd, 2011). The treatment of PV-infected
cells with actinomycin D, an inhibitor of cellular transcription has a negative impact on SG
formation (Piotrowska et al., 2010), suggesting that cellular transcription may be important
for SG assembly during PV infection. More recently, Wu et al., 2014 have demonstrated that
the viral proteinase 2Apro is necessary to trigger SG formation early in infection (3 h.p.i) by
both CVB3 and enterovirus 71 (EV71), suggesting that other picornaviruses might use the
same mechanism to initiate SG responses (Wu et al., 2014). While the exact mechanism of
2Apro-mediated SG formation remains unclear, 2Apro proteinase activity is critical to initiate
SGs formation (Wu et al., 2014). It was proposed that this may be via the ability of 2Apro to
re-localize SRp20, a nuclear-cytoplasmic shuttling splicing factor, from the nucleus to the
cytoplasmic granules (Fitzgerald et al., 2013), or through the cleavage of eIF4G by 2Apro (Wu
et al., 2014). Other studies have also provided evidence for the absence of G3BP1 and eIF4G
from SGs at late stages of CVB3 and EV71 infections (White et al., 2007; Wu et al., 2014).
39
1.6.3 Inhibition of stress granule formation by modulation of eIF2 phosphorylation
Responding to viral entry, mammalian orthoreovirus (MRV), a Reoviridae, induces
SG formation early during infection (6hpi) and this correlates with an increase in eIF2
phosphorylation and reduced cellular translation (Qin et al., 2011). These events may
promote viral uncoating which depends on eIF2 phosphorylation and expression of ATF4, a
transcriptional factor that enhances viral replication when cellular mRNAs localise to SGs
(Smith et al., 2006b). Interestingly, at this stage SGs contain viral core particles but the role
of this is unclear (Qin et al., 2009). Associated with an increase of viral translation, the
disassembly of SGs is observed late in infection (24 hpi) as the replication cycle progresses
independently from eIF2α phosphorylation (Qin et al., 2011). Moreover, the knockdown of
PKR or other eIF2α kinases has no effect on SG formation, indicating that the induction of
SGs during MRV infection may occur by involving two or more eIF2α kinases and through
the contribution of viral proteins (Qin et al., 2009). These findings clearly highlight that
MRV regulates assembly/disassembly of SGs as the replication cycle progresses. The
induction of eIF2α-dependent SGs early during infection and their blockage late in infection
was also shown in mouse embryo fibroblasts (MEF) during Semliki Forest Virus (SFV)
infection (McInerney et al., 2005).
Recently, Ruggieri et al., 2012 developed a live-cell imaging approach to demonstrate
that HCV utilises specific mechanisms to control SG assembly/disassembly throughout the
infection cycle via the oscillation of SG formation, preventing prolonged translation
repression and promoting viral replication. This mechanism depends on DNA damage–
inducible protein 34 (GADD34), the cofactor of phosphatase1 that dephosphorylates eIF2α
and PKR, regulating the phosphorylation status of eIF2α (Ruggieri et al., 2012). Junín virus
(JUNV) is another example of a virus that mediates blockage of SG formation. JUNV
40
infection does not induce SG formation in Vero cells, and the induction of SGs is impaired in
infected cells exposed to sodium arsenite-mediated stress (Linero et al., 2011). Studies in
transfected cells revealed that the inhibition of SGs during JUNV infection is mediated by
two viral proteins, nucleoprotein or N protein and glycoprotein precursor (GPC) which affect
eIF2α phosphorylation (Linero et al., 2011). By contrast, the assembly of SGs was observed
in JUNV- infected cells treated with the eIF4A inhibitor, hippuristanol, which induces SG
formation independently from eIF2α phosphorylation (Linero et al., 2011).
1.6.4 Inhibition of stress granule formation by sequestering or co-opting their
components
The sequestration of the SG protein G3BP1 through its interaction with both NS5B
protein of hepatitis C virus (HCV) and the 5′ end of the HCV minus-strand RNA, late in
infection (48 hpi), causes inhibition of SGs induced earlier in infection via PKR activation
(Yi et al., 2011). This study also demonstrated that G3BP1 co-localizes with HCV replication
complexes (RCs) in cells expressing an HCV replicon. HCV replication was dramatically
reduced by siRNA-mediated knockdown of G3BP1, indicating that RC-associated G3BP1
may be co-opted as a functional protein for viral replication.
The sequestration of G3BP1 to RC during SFV infection via its interaction with the
C-terminal domain of the viral non-structural protein 3 (nsP3) was also found to reduce the
formation of SGs early in infection and inhibit their formation late during infection (Panas et
al., 2012). Another example is human T-cell leukaemia virus type 1 (HTLV-1), a
Retroviridae member which uses a different strategy to block SG formation involving
sequestration of another protein critical for assembly of SGs. This strategy is mediated by
Tax protein which is found in the nucleus under normal conditions but shuttles to the
41
cytoplasm in response to different types of stress (Legros et al., 2011). Within the cytoplasm,
Tax associates with histone deacetylase 6 (HDAC6), a protein involved in SGs assembly
(Kwon et al., 2007), thereby impairing SGs formation (Legros et al., 2011). Cricket paralysis
virus (CrPV), a member of the Dicistroviridae family also blocks SGs formation by
preventing Rox 8 and Rin, the Drosophila homologs of TIA-1 and G3BP1, respectively, from
localising in infected Drosophila S2 cells, even in the presence of stress factors such as heat
shock, arsenite or pateamine A (Khong & Jan, 2011). Other flaviviruses, such as West Nile
virus (WNV) and dengue virus (DV), also have RC colocalizing with TIA-1 and TIAR
proteins via an interaction with the 3′ end of the WNV viral genome thus impairing SGs
assembly (Emara & Brinton, 2007; Li et al., 2002). These viruses take advantage of this
antiviral response, because the lack of these proteins, for example TIAR-depletion in MEF
cells, impairs viral replication (Li et al., 2002). In another flavivirus, Japanese encephalitis
virus (JEV), the interaction between the core structural protein and Caprin-1 (a SGs
component) mediates the inhibition of SGs formation via the recruitment of several SG-
associated proteins, including Caprin-1, G3BP1 and USP10, thereby promoting viral
propagation (Katoh et al., 2013).
Rotavirus has evolved a mechanism to block SGs despite eIF2 phosphorylation
during the early stage of infection (Rojas et al., 2010). Montero et al., 2008 suggested that
this overcoming of eIF2 phosphorylation-mediated translation arrest may be due to the
relocation of PABP (a component of SGs) from the cytoplasm to the nucleus thereby
preventing the formation of SGs (Montero et al., 2008).
In the case of human immunodeficiency virus 1 (HIV-1) infection, Staufen1 is
recruited to RNPs containing genomic RNA via its interaction with Gag inhibiting SGs
formation (Abrahamyan et al., 2010). The mechanism by which HIV-1 blocks SGs assembly
42
also involves the interaction of viral Gag protein with eukaryotic elongation factor 2 (eEF2)
and this process occurs irrespective of eIF2α phosphorylation or overexpression of G3BP1 or
TIAR, which generally trigger SGs formation (Valiente-Echeverria et al., 2014). By contrast
with HIV-1, HIV-2 expression induces stress granule assembly. Moreover, HIV-2 genomic
RNA associates with TIAR to form a TIAR-HIV-2 ribonucleoprotein (TH2RNP), which is
diffused in the cytoplasm or accumulated in SGs in the absence of active translation (Soto-
Rifo et al., 2014). Cycloheximide or puromycin treatment lead to either complete localization
of the TH2RNP complex diffused throughout the cytoplasm, or the incorporation of this
complex in enlarged SGs indicating the TH2RNP complex is in equilibrium between
polysomes and SGs. Nevertheless, the formation of TH2RNP complex does not require
Gag/gRNA binding, but the level of the Gag polyprotein could regulate translation and the
cytoplasmic localization of this complex. A low concentration of Gag promotes the
translation of gRNA and diffused localisation of the gRNA and TIAR in the cytoplasm,
whereas a high level of Gag inhibits the translation of gRNA leading to accumulation of
gRNA/TIAR in SGs. The ability of the gRNA and Gag to accumulate in SGs may reflect
their role as sites where the switch from viral translation to packaging occurs (Soto-Rifo et
al., 2014).
Theiler’s murine encephalomyelitis virus (TMEV), a picornavirus, inhibits SGs
assembly induced by sodium arsenite or thapsigargin treatment in a process mediated by the
leader (L) protein (Borghese & Michiels, 2011). TMEV viral L protein inhibits the host
immune response by disrupting IFN and chemokine production thereby enhancing viral
persistence (van Pesch et al., 2001). Interestingly, the L protein prevents SG formation
independently from its inhibition of interferon production. This raised the possibility that the
inhibition of SGs formation is due to the role of the L protein in altering nucleocytoplasmic
trafficking factors involved in SG assembly (Borghese & Michiels, 2011). NS1 protein-
43
mediated PKR inhibition is essential to block SG formation at a late stage of influenza A
virus (IAV) infection (Khaperskyy et al., 2012). Khaperskyy et al., 2014 demonstrated that
two other proteins can inhibit SGs formation during IAV infection and that the nucleoprotein
(NP) and host-shutoff protein polymerase-acidic protein-X (PA-X) inhibit SGs formation
independently from eIF2α phosphorylation. NP and PA-X action are concomitant with a
dramatic decrease and increase of PABP in the cytoplasm and nucleus respectively
(Khaperskyy et al., 2014). Furthermore, the interaction between NS1 and RNA-associated
protein 55 (RAP55) also impairs SGs formation (Mok et al., 2012).
1.6.5 Viral replication mechanism that benefits from stress granule formation
Some viruses can replicate and cope with the presence of SGs during infection. For
example, despite some effect, the mouse hepatitis coronavirus (MHE) replicative cycle
continues during infection while SGs are stimulated through eIF2 phosphorylation (Raaben
et al., 2007). The replication of respiratory syncytial virus (RSV) is also completed despite
the presence of SGs induced via PKR-mediated mechanisms (Lindquist et al., 2011).
In the case of negative-stranded RNA viruses, Vesicular stomatitis virus (VSV)
infection induces unconventional SGs because these granules do not contain some of the
bona fide SG markers, such as eIF3 or eIF4A, or the processing body (PB) markers, Dcp1a
and so-called SG-like structures which contain poly-C binding protein (PCBP2), TIA1 and
TIAR. Interestingly, VSV SG-like structures contain the viral replicative proteins (VSV N
protein) and viral RNA. Moreover, viral replication and protein synthesis as well as the
presence of TIA-1 are necessary for forming the SG-like structures during VSV infection
(Dinh et al., 2013). In this study, the formation of SG-like structures had no significant effect
44
on viral protein synthesis despite increased levels of phosphorylated eIF2 at different times
post-infection.
All the examples mentioned above are RNA viruses; few studies have investigated the
relationship between SGs formation and DNA viruses (Esclatine et al., 2004; Isler et al.,
2005). For example, herpes simplex virus-1 (HSV-1), SGs are not observed during infection
(Esclatine et al., 2004). However, the induction of SGs using sodium arsenite, a classical
inducer of oxidative stress is also inhibited in HSV-2 infected cells, whereas the SGs
containing G3BP1 and PABP but not TIA-1 were observed in infected cells that treated with
Pat A (Finnen et al., 2012). This may be due to an interaction of virion host shutoff (Vhs)
protein (HSV virion component) with a key component of SGs such as TTP (Dauber et al.,
2011; Esclatine et al., 2004; Finnen et al., 2014). The infection with vhs-deficient mutants of
HSV-1 or -2 induces SGs formation late in infection, these granules contain core SG
components such as TIA-1, TIAR and G3BP1 as well as the viral immediate early protein
ICP27 and the viral serine/threonine kinase Us3 (Esclatine et al., 2004; Finnen et al., 2014).
45
1.7 Virus/P-bodies interaction
Compared with SGs, there are only a few studies describing the relationship between
PBs and viruses, most of which are related to positive-stranded RNA viruses (Pattnaik &
Dinh, 2013; Tsai & Lloyd, 2014). In addition to the role of PBs in suppressing certain viral
life cycles, other mechanisms for the disruption of PBs during viral infection or viral co-
opting of PBs components have been described (Lloyd, 2013) (Figure 1.6).
In addition to 3CPro cleaving G3BP1 during PV infection, Dougherty et al, 2011 also
proposed a role for this enzyme in the rapid disruption of PBs 2-4h after PV and CVB3
infection through the cleavage or degradation of key components of PBs such as Xrn1, Dcp1a
and Pan3, suggesting an inhibition of the de-adenylation which may be required for viral
replication (Dougherty et al., 2011). In addition, WNV and DV impair PB assembly at a late
stage of infection (24-36 hpi) and the reduction of PBs was shown to be concomitant with the
relocation of TIA-1 and decreased eIF2 phosphorylation level in cells exposed to oxidative
stress via unidentified mechanisms (Emara & Brinton, 2007). Later, Chahar et al., 2013
reported that WNV infection causes the recruitment of many PB components including
Lsm1, GW182, DDX3, DDX6 and Xrn1 that localise with viral non-structural protein 3
(NS3) at viral replication sites. Some of these proteins have been demonstrated to be positive
regulators of viral replication as their siRNA- mediated knockdown impairs viral replication
(Chahar et al., 2013). The progressive decrease in PBs was also observed at a late stage
during HCV infection. DDX6 or G3BP1 were found to colocalise with HCV core 48 h after
infection (Ariumi et al., 2011), indicating that the HCV hijacks the P-body and the stress
granule components to regulate HCV RNA replication and translation. Supporting this,
knockdown of DDX3 reduced HCV replication. Other PB constituents including Ago2,
Lsm1, PatL1, Ge1, and miR122 are required for HCV replication as their knockdown had a
46
negative impact on viral replication (Ariumi et al., 2011; Berezhna et al., 2011; Pager et al.,
2013; Thibault et al., 2013).
Recently, using hepatocytes from human livers of patients infected with different
genotypes of HCV, Perez-Vilano et al., 2015 were also able to demonstrate disruption of P-
body formation in vivo (Perez-Vilaro et al., 2014). They also demonstrated that this inhibition
was relieved upon HCV elimination using antiviral therapy. Interestingly, they also provided
evidence for a high heterogeneity in the composition and shape of PBs formed in uninfected
hepatocytes. This clearly demonstrates a link between PBs formation and a key pathogenic
condition.
Other examples of viruses that co-opt components of PBs to enhance their replication
are HIV and MLV, as the replication of both viruses is affected by the depletion of Mov10, a
PBs protein (Burdick et al., 2010; Furtak et al., 2010). Moreover, HIV virion assembly is
reduced by knockdown of DDX6 and Ago2 which both localise with HIV Gag protein at PBs
assembly sites (Reed et al., 2012). Among negative-sense RNA viruses, NS1 viral protein
mediates the disruption of PBs during influenza virus infection via its interaction with Rap55,
preventing the sequestration of the viral NP and NP-associated RNAs in PBs thereby
promoting viral replication (Mok et al., 2012). The Hantavirus nucleocapsid protein binds to
the 5′ cap of cellular mRNAs and protects the 5′ extremity from decapping and degradation
during accumulation in PBs, and is involved in initiation of viral transcription (Mir et al.,
2008).
Less is known about the relationship between PBs and DNA viruses. However, the
adenovirus replication cycle causes PB reduction by altering PB components through a
specific interaction between viral proteins and DDX6. This association causes the
redistribution of several PB constituents to aggresomes (Greer et al., 2011).
47
Figure 1.6 Mechanisms of P-bodies disruption by viruses. Viruses evolved many
mechanisms to disrupt or prevent PBs. (A) HCV, flavivirus, and adenovirus can induce the
sequestration of different PBs components to their replication complex (HCV and flavivirus)
or to agressome (adenovirus), disrupting PBs assembly. (B) Viral protease from picornavirus
cleavage an important nucleating-PB proteins including Dcp1, Xrn1 and Pan3 disrupting PBs
assembly. (C) Interactions between specific viral proteins and PB components also prevent
formation of PBs.
48
1.8 Caliciviruses
The Caliciviridae were adopted as a family of viruses in the third report of the
International Committee on Taxonomy of Viruses (ICTV) in 1979, and before this
caliciviruses were considered as part of the Picornaviridae family (Green et al., 2000).
According to the last 2011 report of ICTV, the caliciviruses can be classified into five genera
(shown in Table 1.3) based on their genomic organisation and sequence similarities. These
genera are Lagovirus, Vesivirus, Norovirus, Sapovirus and Nebovirus (Green, 2013;
International Committee on Taxonomy of Viruses, http://ictvonline.org). Five additional
genera have been proposed including "Salovirus" (Mikalsen et al., 2014), "Bavovirus"(Wolf
et al., 2012), "Nacovirus" (Day et al., 2010; Wolf et al., 2012), "Recovirus" (Farkas et al.,
2014; Farkas et al., 2008) and "Valovirus" (L'Homme et al., 2009), and several Caliciviridae
remain unclassified (including 18 species; http://www.caliciviridae.com).
Caliciviruses are small, non-enveloped, positive-sense single-stranded RNA viruses.
Their genomes range between 7.3 to 8.5 kb in length, containing 2 to 4 open reading frames
(ORFs), and are surrounded by a small icosahedral capsid of 27 to 40 nm in diameter
(Rohayem et al., 2010; Thackray et al., 2007). The name calicivirus originates from the Latin
word for cup (“calyx”) because of the cup-shaped indentations observed on the virion using
electron microscopy (Thiel & Konig, 1999) (Figure 1.7)
The Caliciviridae family includes viruses that cause gastrointestinal disease in human
hosts and a broad spectrum of diseases in a wide variety of animal hosts (Glass et al., 2000b;
Rohayem et al., 2010) leading to significant economic impact both to the UK National Health
Service (NHS) and businesses in the case of human disease, and to the agricultural industry
and farming for animals.
49
Table 1.3 Taxonomy of the Caliciviridae family (ICTV, 2011); Caliciviruses taxonomy
browser, 2014)
Genus Species Viruses
Vesivirus
(2 Species - 46 complete
genomes)
Feline calicivirus
(711 strains - 32 complete genomes) Feline calicivirus serotypes
Vesicular exanthema of swine virus
(99 strains - 6 complete genomes)
Vesicular exanthema of swine virus serotypes, bovine calicivirus,
cetacean calicivirus, primate calicivirus, reptile calicivirus, San Miguel
sea lion virus serotypes
Sapovirus
(1 Species - 40 complete
genomes)
Sapporo virus
(2162 strains - 38 complete genomes)
Porcine enteric calicivirus, Sapporo virus, Houston virus, Manchester
virus
Norovirus
(1 Species - 688 complete
genomes)
Norwalk virus
(17729 strains - 624 complete
genomes)
Murine norovirus, Lordsdale norovirus, Norwalk virus, Snow
Mountain norovirus, Southampton norovirus
Lagovirus
(2 Species - 41 complete
genomes)
European brown hare syndrome virus
(82 strains - 4 complete genomes) European brown hare syndrome virus serotypes
Rabbit haemorrhagic disease virus
(547 strains - 35 complete genomes) Rabbit haemorrhagic disease virus serotypes
Nebovirus
(1 Species - 5 complete
genomes)
Newbury-1 virus
(15 strains - 4 complete genomes) Newbury-1 virus, Nebraska virus
Unclassified Caliciviridae
(18 Species - 20 complete
genomes)
(18 Species - 20 complete genomes)
Calicivirus strain HP167-KOR, Calicivirus strain MA120-KOR,
Calicivirus strain MA129-KOR, Calicivirus strain MA137-KOR,
Calicivirus strain MA16-KOR, Calicivirus strain MA64-KOR,
Calicivirus strain MA76-KOR, Calicivirus strain MA176-KOR,
Calicivirus strain MA216-KOR, Calicivirus strain MA236-KOR,
Calicivirus strain MA360-KOR, Calicivirus strain MA56-KOR
Calicivirus strain MA91-KOR, Calicivirus strain MA95-KOR,
Calicivirus strain MA99-KOR, Calicivirus strain SA181-KOR,
Calicivirus strain SA68-KOR, Calicivirus strain MA129-KOR,
Recombinant virus 6918 VP60-T2
50
Figure 1.7 Crystal structures of Norwalk Virus, MNV-1 and FCV by cryo-electron
microscopy from (Bhella et al., 2008; Katpally et al., 2008; Prasad et al., 1999). (A)
Structure of HuNoV (Norwalk Virus); (B) Structure of MNV-1; (C) Structure of FCV. The
cup-shaped indentations characteristics of caliciviruses are visible in green in A, blue in B,
and pink in C.
51
1.8.1 Lagoviruses (LaVs)
The genus Lagovirus consists of two species, Rabbit Haemorrhagic disease virus
(RHDV) and European Brown Hare Syndrome Virus (EBHSV) (ICTV, 2013). RHDV was
first detected in China in 1984 and associated with the death of 14 million domesticated
angora rabbits within nine months. The virus then rapidly spread worldwide via Italy to the
rest of Europe and is now present as an endemic agent in many countries (Abrantes et al.,
2012a; Abrantes et al., 2012b). All pathogenic strains are classified within a single serotype.
RHDV causes high mortality in both domestic and wild adult animals (Abrantes et al., 2012a;
McIntosh et al., 2007). The disease is characterised by acute necrotising hepatitis, but other
organs, such as lungs, heart, and kidney may also be affected (Abrantes et al., 2012b). The
transmission of RHDV is mainly via direct contact with infected rabbits through the oral
route and skin lesion (Merchan et al., 2011), or indirectly via contaminated food, bedding,
water, clothes and veterinary equipment (Cooke, 2002). Despite the similarities between
RHDV and EBHSV in clinical signs, pathological and histopathological alterations, mortality
rates, virion morphology and antigenicity, these viruses infect different species (Abrantes et
al., 2012b). EBHSV was first detected in Sweden in 1980 (Gavier-Widen & Morner, 1991),
and is the causative agent of European brown hare syndrome in Lepus europaeus and Lepus
timidus (Lopes et al., 2014). Subsequently, the virus was recognised in many European
countries (Paci et al., 2011) and now is considered endemic in all European countries (Chiari
et al., 2014). EBHSV is a highly contagious disease and transmission occurs via direct
contact or through the oral-fecal route (Frolich et al., 2007).
52
1.8.2 Vesiviruses (VeVs)
The Vesivirus genus is represented by two species, Vesicular exanthema of swine
virus (VESV), which consists of many serotypes and feline calicivirus (FCV) with a single
serotype. In addition, a number of vesiviruses remain unclassified (ICTV, 2013). VESV was
first observed in 1932 in California, USA, associated with vesicular disease in swine. From
1952 the disease spread through most USA states leading to massive economic losses (Smith
et al., 2006a). Later in 1972, related viruses were isolated from sea lions on San Miguel
Island, California and named as San Miguel Sea lion virus (SMSV), causing vesicular disease
when transferred to swine (Smith et al., 1973). This led to the suggestion that all cases of
VESV in USA between 1932 and 1959 were caused by marine origin serotypes (Reid et al.,
2007; Smith et al., 2006a). Importantly, Vesivirus of marine origins have been found to infect
humans (Smith et al., 2006a). In addition to the SMSV serotype, VESV species have several
serotypes infecting a wide range of hosts such as fish, reptiles, primates and others animals
(Smith et al., 2006a; Wellehan et al., 2010).
A second species of Vesivirus is feline calicivirus (FCV), a highly infectious pathogen
that infects domestic and wild cats, and is responsible for acute oral and upper respiratory
tract disease with a range of clinical signs which depend on the virulence of FCV strains
(Geissler et al., 1997; Radford et al., 2009). Some strains are associated with chronic
stomatitis and limping syndrome (Radford et al., 2009). The routes of transmission are nasal,
oral or conjunctival routes, and following infection, the replication occurs in the oropharynx.
The infection starts with a necrosis in epithelial cells which subsequently leads to ulcer
formation (Radford et al., 2009). FCV can cause pneumonia and lameness (Dawson et al.,
1994). Recently, highly virulent strains of FCV with high mortality (up to 67%) have been
recognised in the USA and Europe and cause FCV-associated virulent systemic disease
(FCV-VSD). They are responsible for outbreaks of severe and acute systemic disease which
53
differs from the more usual FCV disease (Geissler et al., 1997; Ossiboff et al., 2007;
Pesavento et al., 2004; Radford et al., 2007). These strains are characterised by a new range
of clinical signs including systemic inflammatory response syndrome, jaundice, ulcerative
dermatitis and oedema that can lead to death (Coyne et al., 2012; Pesavento et al., 2004;
Radford et al., 2009). Vaccination can prevent the disease but does not prevent the infection
(Coyne et al., 2006; Southerden & Gorrel, 2007). To explain this virulence, it was found that
the most variable region of FCV is located in the surface of the capsid protein, representing
the main target for the immune response (Radford et al., 2009). The evolutionary rate of this
region is up to 1.3x10-2 to 2.6x10-2 substitution per nucleotide per year, the highest
evolutionary rate reported compared with any virus (Coyne et al., 2012). In addition, FCV
displays resistance to common disinfectants and can persist in the environment for 1 month
(Radford et al., 2009). Importantly, FCV is used as a model for the study of the molecular
biology of human norovirus (Vashist et al., 2009).
1.8.3 Sapoviruses (SaVs)
The Sapovirus genus consists of a single species divided into five genogroups (GI-V),
and subdivided into a number of genotypes based on the capsid gene sequence (Hansman et
al., 2007; Oka et al., 2012). Saporoviruses are responsible for enteric diseases in human and
animal hosts (Hansman et al., 2007; L'Homme et al., 2010). To date, only genogroup GIII is
associated with animal disease especially in porcine species (Di Bartolo et al., 2014; Farkas
et al., 2004; Hansman et al., 2007; L'Homme et al., 2010; Pang et al., 2009). The other
genogroups (GI, GII, GIV and GV) have been reported as causative agents of human acute
gastroenteritis worldwide (Bucardo et al., 2014; Hansman et al., 2007; Nidaira et al., 2014;
Svraka et al., 2010; Usuku & Kumazaki, 2014; Wang et al., 2014). They can affect all age
54
groups but are more common in children (Dey et al., 2012; Hansman et al., 2007; Lee et al.,
2012; Sakai et al., 2001; Usuku & Kumazaki, 2014; Wu et al., 2008). Several reports have
also reported outbreaks of SaV in adults (Lee et al., 2012; Nidaira et al., 2014; Wang et al.,
2014). The first report of an SaV outbreak was by Chiba et al., 1977 using electron
microscopy of stool specimens from infants suffering from acute gastroenteritis in an
orphanage in Sapporo, Japan (Chiba et al., 1979). The genogroups GI/II have been reported
as the most frequent strains associated with outbreaks of gastroenteritis worldwide (Hansman
et al., 2007; Lee et al., 2012; Miyoshi et al., 2010; Nidaira et al., 2014; Ootsuka et al., 2009;
Usuku & Kumazaki, 2014; Wang et al., 2014).
Porcine SaV (PSaV) has been detected in symptomatic or diarrheic pigs (Collins et
al., 2009; Di Bartolo et al., 2014; L'Homme et al., 2009; Scheuer et al., 2013) and recently
numbers of novel animal SaV strains related to GIII and human strains have been identified
in mink and swine (Guo et al., 2001; Martella et al., 2008; Wang et al., 2005; Yin et al.,
2006). In addition, PSaV can be propagated in tissue culture, providing a good model for
understanding the molecular aspects of human SaV (Chang et al., 2002; Hosmillo et al.,
2014). More recently, SaV GI viruses have been detected in faecal samples from
chimpanzees living in close contact with humans in the Democratic Republic of Congo
(Mombo et al., 2014). Human SaV strains have been reported in food and water in areas
surrounding populations that have suffered from SaV outbreaks (Iizuka et al., 2010;
Kobayashi et al., 2012; Murray et al., 2013; Sano et al., 2011). Therefore, it is likely that the
transmission routes are faecal-oral as well as person-to-person (Sala et al., 2014). The close
genetic relationship between the SaV identified recently in both animals and humans, as well
as in food and water, may trigger further research to address the zoonotic origins of SaV
infections (Bank-Wolf et al., 2010).
55
1.8.4 Neboviruses (NeVs)
The most recent genus of the Caliciviridae family to be established, Nebovirus
contains a single species, Newbury-1 virus (Carstens, 2010). It comprises several strains such
as Newbury “agent”-1, first described in the UK in 1976 and Nebraska virus first detected in
Nebraska, USA in 1980 (Carstens, 2010; Oliver et al., 2006; Smiley et al., 2002). So far, all
strains described in this genus are bovine enteric viruses (Smiley et al., 2002) ICTV, 2013).
Recently, the frequent detection of an association between bovine caliciviruses such as
Nebovirus and Bovine norovirus (BNoV) with calf diarrhoea suggested that these viruses are
a significant cause of endemic diarrheal disease in calves (Cho et al., 2013; Kaplon et al.,
2011; Park et al., 2007). As the majority of caliciviruses, Nebovirus cannot be propagated in
cell culture (Carstens, 2010).
1.9 Noroviruses (NoVs)
NoVs are the most important causative agent of viral gastroenteritis outbreaks in
humans worldwide, but have also been detected in some animal hosts (Hall et al., 2011;
Ahmed et al., 2014; Green et al., 2000). This genus contains only one species, based on
phylogenetic analysis of their capsid gene (and recently of ORF1 sequence), which can be
divided into six genogroups (GI-GVI) containing several genotypes (Caddy et al., 2014;
Green et al., 2000; Kroneman et al., 2013; Zheng et al., 2006)(Figure 1.8). Animal NoVs
belong to genogroups GIII (bovine NoV), GV (murine NoV), GII (porcine NoV) and GIV
(lion NoV) (Caddy et al., 2014; Martella et al., 2007; Scipioni et al., 2008; Wang et al., 2005;
Zheng et al., 2006), whereas human NoVs fall in GI, II and IV (Atmar, 2010; Caddy et al.,
2014; Green et al., 2000; Zheng et al., 2006). Genetic relationships between human and
animal NoVs may reflect a potential zoonotic transmission (Scipioni et al., 2008).
56
Figure 1.8 Phylogenetic analysis of the capsid amino acid sequence of Norovirus
strains. Norovirus can be classified into six genogroups (GI-GVI) based on their capsid
gene and ORF1 sequence containing several genotypes as indicated. Genotype GII.4 has
seven different variants of genotypes as boxed. (Modified from (Glass et al., 2009;
Kroneman et al., 2013; Patel et al., 2009; Ramani et al., 2014).
57
1.9.1 Animal noroviruses
To date, animal noroviruses have been detected in cattle, pigs (Karst et al., 2003;
Oliver et al., 2003; Wang et al., 2005; Wolf et al., 2009), dogs and rats (Martella et al., 2007;
Pinto et al., 2012; Tse et al., 2012), lion cubs (Martella et al., 2007) and mice (Karst et al.,
2003), representing infection by bovine, porcine, canine, feline and murine noroviruses.
Diarrhoea is the common clinical sign reported in calves infected with NoV but is not
reported in infected pigs or mice (Dastjerdi et al., 1999; Hsu et al., 2007; Wang et al., 2007;
Wang et al., 2006)
1.9.2 Murine norovirus-1 (MNV-1)
MNV-1 was first discovered in 2003 in the USA and associated with lethal infection
of laboratory mice (Karst et al., 2003). Many studies have since then reported MNV
infections in laboratory mice in different countries (Hsu et al., 2005; Mahler & Kohl, 2009;
Ohsugi et al., 2013). Although MNV-1 can infect healthy mice it does not result in any
disease (Thackray et al., 2007). Mice lacking recombination-activation gene 2 (RAG2) and
activator of transcription 1 (STAT1), two factors involve in the immune responses, are very
sensitive to MNV-1, which can replicate within lymphoid tissues and display many clinical
signs leading to death (Hsu et al., 2006; Karst et al., 2003). This indicates that the innate
immune system components have a very strong effect on MNV-1 infection. To date, 15
distinct MNV strains have been isolated such as MNV-2, 3 and 4 (Hsu et al., 2006; Thackray
et al., 2007). These strains have different biological properties or pathogenic patterns but all
belong to one MNV serotype (Goto et al., 2009; Hsu et al., 2006; Thackray et al., 2007).
Transmission of MNV is mainly via the faecal-oral route especially through contaminated
bedding (Manuel et al., 2008). It has been demonstrated that MNV strains are able to
58
replicate and persist in different cells and tissues such as macrophages and dendritic cells
(Wobus et al., 2004) as well as the small intestine, caecum, mesenteric lymph nodes and the
spleen of both immune-competent and immune-deficient mice (Hsu et al., 2006; Thackray et
al., 2007). Other studies have reported that MNV circulates in wild rodents (Farkas et al.,
2012; Smith et al., 2012). MNV is the only norovirus that replicates in cultured cells, mainly
macrophage cell lines; MNV-1 can be propagated in RAW 264.7, BV-2 and TIB cell lines
(Cox et al., 2009). Because of these properties, and the genetic relationship to human NoVs,
MNV provides an excellent model to study the translation and replication mechanisms of
Human NoVs (Thorne & Goodfellow, 2014; Wobus et al., 2006).
1.9.3 Human noroviruses (HuNoVs)
In recent years, human noroviruses have been recognised as a cause of gastroenteritis
disease worldwide and are commonly associated with acute gastroenteritis outbreaks (Glass
et al., 2009; Patel et al., 2009). Outbreaks mainly occur in enclosed settings such as hospitals,
schools, nursing homes as well as cruise ships or military barracks (Bert et al., 2014;
Donaldson et al., 2008; Godoy et al., 2014; Hansen et al., 2007; Iturriza-Gomara & Lopman,
2014; Simon et al., 2006; Widdowson et al., 2004; Xue et al., 2014). HuNoVs are responsible
for the winter vomiting disease (also named stomach flu or gastric flu) which happens more
commonly in winter and affects people of all ages but has a more deleterious effect on
children, elderly and immune-compromised people (Estes et al., 2006; Green, 2014; Payne et
al., 2013; Rackoff et al., 2013). Each year, HuNoVs are responsible for 50% of all
gastroenteritis outbreaks (over 267,000,000 infections) (Donaldson et al., 2008) and
approximately 200,000 deaths especially in children under 5 years of age and the elderly (˃65
years) (Hall et al., 2013; Patel et al., 2008).
59
Winter vomiting disease was first recognised in 1929 by Dr Zahorsky and named as a
“Hyperemesis hemis’’ or winter vomiting disease, but the causative agent for this disease was
unknown. 43 years later, Albert Kapikian investigated stool samples from a winter vomiting
disease outbreak that occurred in an elementary school in Norwalk, Ohio, USA in 1968 and
identified a virus, naming this virus Norwalk virus (a prototype of noroviruses). It was
established as the causative agent using immune-electron microscopy (IEM) to examine stool
material from adult volunteers administered with faecal filtrate from this gastroenteritis
outbreak (Kapikian et al., 1972), later the name was changed to “small round structure virus”
(Green et al., 2000). HuNoVs are found mainly in GI and GII and few GIV norovirus
genogroups (Caddy et al., 2014; Kroneman et al., 2013; Rackoff et al., 2013; Siebenga et al.,
2009; Zheng et al., 2006). The symptoms vary depending on many factors such as age but the
most common symptoms are vomiting especially in infants, and diarrhoea as well as nausea,
abdominal pain, abdominal cramps, anorexia, myalgia and low fever (Atmar & Estes, 2006;
Patel et al., 2009). Recently, the numbers of HuNoV infection have increased and HuNoVs
cause 18% of all cases of gastroenteritis and are responsible for more than 50% of all
gastroenteritis outbreaks worldwide and between 75/90% of nonbacterial gastroenteritis
outbreaks in developing countries (Ahmed et al., 2014; Hall et al., 2014; Patel et al., 2008;
Scallan et al., 2011; Siebenga et al., 2009). In the United States, the number of HuNoV cases
is between 19 and 23 million each year, leading to 56,000-71,000 hospitalisations and 570-
800 deaths which cost the government about $777 million every year (Hall et al., 2013).
Overall the cost associated with foodborne outbreaks is $2 billion (Glass et al., 2009; Hall et
al., 2013; Hoffmann et al., 2012). In the United Kingdom, annual expenses associated with
HuNoV infection are estimated to be over £100 million (Lopman et al., 2004).
The most common genotype associated with the majority (~80%) of HuNoV
gastroenteritis including food-borne outbreaks is GII.4 whereas GI which is more stable in
60
water is responsible for water-borne outbreaks (Matthews et al., 2012; Patel et al., 2008;
Rackoff et al., 2013). Worldwide, 55 to 85% of cases and outbreaks from 1990s to 2013,
which included four pandemics, were caused by seven different variants of genotypes GII.4
including: the US 1995/96 variant in 1996 (White et al., 2002), the Farmington Hills variant
in 2002 (Widdowson et al., 2004), the Hunter variant in 2004 (Bull et al., 2006), the 2006a,
2006b variants in 2007-2008 (Eden et al., 2010), and the New Orleans variant from 2009 to
2012 (Yen et al., 2011). More recently, a new variant of genotype GII.4 of HuNoVs (called
Sydney2012) was reported in New South Wales, Australia (van Beek et al., 2013). This then
spread around the world to replace the previous variant of genotype GII.4 (New Orleans
2009) as a commonly detected HuNoV strain in many countries (Allen et al., 2014; da Silva
et al., 2013; Fonager et al., 2013; Leshem et al., 2013; Shen et al., 2013; van Beek et al.,
2013; Vega et al., 2014). No significant differences in the number of outbreaks between
Sydney 2012 and New Orleans 2009 strains exist, but the new strain is associated with
distinct genetic changes in the P2 component of the capsid protein VP1 sequence compared
with that of the New Orleans 2009 strain (Allen et al., 2014; Debbink et al., 2013). These
changes to six amino acid positions, including 294, 310, 359, 368, 373 and 396, play an
important role in defining histo-blood group antigens (HBGAs) leading to the antigenic
variation responsible for increasing fitness of Sydney 2012 strains. These antigenic variations
play a critical role for the emergence of a novel strain by promoting viral escape from host
immunity (Allen et al., 2014; Debbink et al., 2012).
Typically, a new variant emerges every 2-3 years to replace the previous dominant
variant of genotype (Siebenga et al., 2007, Desai et al., 2012, Gastanaduy et al., 2013). Most
variants usually emerge within the GII.4 genotype but also among GII.2 and GI.3 (Zheng et
al., 2006, Eden et al., 2013). Four factors can influence the evolution rate of NoVs; these are
61
host receptor recognition, sequence space, duration of population immunity, and replication
kinetics (Bull & White, 2011).
1.9.4 Clinical signs
The incubation period of HuNoV is usually 24-48 hrs and the symptoms vary
depending on factors such as age and the infectious dose (Kaplan et al., 1982; Karst, 2010;
Murata et al., 2007; Patel et al., 2009). The illness is commonly characterised by nausea,
vomiting (more common in children below 1 year) and watery diarrhoea (more common in
adults) as well as other clinical features including anorexia and in some cases abdominal
cramps and pain, fever, headache, chills and myalgia (Deng et al., 2007; Kaplan et al., 1982;
Karst, 2010). The duration of illness is usually 2 to 4 days, but can be longer in nosocomial
outbreaks (4-6 days) (Kaplan et al., 1982; Lee et al., 2013; Lopman et al., 2009; Murata et
al., 2007; Rockx et al., 2002; Sukhrie et al., 2011) lasting up to six weeks in infants (Patel et
al., 2009).
Although, the illness is self-limiting in the majority of cases, there is increasing
evidence that norovirus may lead to post infection disorders leading to death in the elderly
and immunocompromised patients (Harris et al., 2008; Rondy et al., 2011; Sukhrie et al.,
2010). In elderly patients mortality can be associated with dehydration (Harris et al., 2008;
van Asten et al., 2011). Studies have recognised chronic infection in immunocompromised
patients (Bok & Green, 2012; Ronchetti et al., 2014; van Asten et al., 2011). HuNoV is also
associated with other important clinical diseases including necrotizing enterocolitis (Stuart et
al., 2010; Turcios-Ruiz et al., 2008) and afebrile seizure in infants (Chan et al., 2011), as well
as pneumatosis intestinalis (Kim et al., 2011) and Crohn’s disease (Cadwell et al., 2010). In
the UK, this leads to around 80 deaths annually among elderly people (older than 64 years)
62
(Harris et al., 2008). In addition, according to the WHO estimations, approximately 200,000
deaths per year occur in children below 5 years of age in developing countries (Patel et al.,
2008).
Despite a short duration of virus infection, prolonged viral shedding is reported for 3-
8 weeks following infection with high viral loads (up to 1012 NoV copies/g faeces) in
symptomatic or asymptomatic patients after recovery (Atmar et al., 2008; Rockx et al., 2002)
shedding can also continue over two years in immunocompromised patients and transplant
patients (Bok & Green, 2012). Moreover, in some cases of paediatric oncology patient the
virus shedding continues for over a year (Simon et al., 2006).
1.9.5 Transmission
Three general routes are involved in HuNoV transmission including; person-to-person
transmission directly via the faecal-oral route (consumption of food or aerosolized vomitus)
or indirectly through contaminated surfaces or fomites. The second route is food-borne
transmission via an infected food handler or food distribution system contaminated with
human waste. Finally, contaminated water is a vehicle for HuNoV outbreaks (Harris et al.,
2014; Isakbaeva et al., 2005; Yoder et al., 2008; Yoder et al., 2004). The transmission of
HuNoV is facilitated by several factors including low virus infectious dose, 8-1000 virus
particles, prolonged viral shedding, environmental stability, genetic diversity and short term
immunity which all play a critical role during HuNoV outbreaks (Debbink et al., 2013;
Duizer et al., 2004a; Hall, 2012; Hansman et al., 2006; Kirkwood & Streitberg, 2008;
Siebenga et al., 2008; Simmons et al., 2013; Teunis et al., 2008; Zheng et al., 2006). The
majority of HuNoV outbreaks occur in closed and semi-closed settings such as schools,
restaurants, cruise ships, nursing homes, and hospitals. In addition to the virus characteristics:
63
low infectious dose, the short incubation period, high viral load in stool and vomitus as well
as environmental stability; the transmission of NoVs in these closed settings is dependent on
other conditions such as factors associated with patients, visitors and staff behaviour (Harris
et al., 2014; Iturriza-Gomara & Lopman, 2014; Lopman et al., 2012; Patel et al., 2009).
1.10 Model systems for the study of human norovirus biology
Since its discovery in 1972, our knowledge of HuNoVs biology, pathogenesis,
immunity, translation and replication remains incomplete due to the lack of a suitable tissue
culture system or any small animal model to propagate and study these viruses (Karst et al.,
2014a; Karst et al., 2014b; Thorne & Goodfellow, 2014). So far, several attempts have been
made to propagate HuNoV in different types of tissue or cell line as well as many animal
models using a variety of methods and techniques, but with very limited success (Cheetham
et al., 2006; Duizer et al., 2004b; Papafragkou et al., 2013; Rockx et al., 2005; Straub et al.,
2011; Straub et al., 2007; Subekti et al., 2002; Takanashi et al., 2014; Wobus et al., 2006).
This impairs the efforts to design and develop drugs or vaccines against these viruses (Karst,
2010; Rocha-Pereira et al., 2014; Rohayem et al., 2010; Thorne & Goodfellow, 2014). Early
attempts used human volunteers to understand some aspects of the NoV pathogenicity as well
as protective immunity (Dolin et al., 1971; Green et al., 1993). However, there are many
limitation for using humans as a model system including difficulty and cost, and
biocontainment requirement for this type of virus (type B pathogen) (Vashist et al., 2009).
Many studies have attempted to infect non-human hosts with HuNoV using different animals
previously reported as susceptible to enteric caliciviruses such as pigs, monkeys and calves
(Vashist et al., 2009). These attempts have largely been unsuccessful, apart from some
positive results in chimpanzees, and macaques (Bok et al., 2011; Subekti et al., 2002; Wyatt
64
et al., 1978). Limited virus replication has been reported in Rhesus macaques (Rockx et al.,
2005) and in pigtail macaque (Subekti et al., 2002) infected with HuNoV. In addition, viral
replication also has been reported to occur in chimpanzees infected with HuNoV, but this led
to different clinical symptoms from those that occur in humans (Bok et al., 2011; Wyatt et
al., 1978). GII of HuNoV replication also can occur in gnotobiotic pigs and calves, leading to
viremia (Cheetham et al., 2006; Souza et al., 2008) and the majority of infected pigs develop
mild diarrhoea (Cheetham et al., 2006). Recently, (Taube et al., 2013) reported limited
replication of human NoVs using Rag-/-γc-/- BALBc deficient mice as a model, but the
infected mice did not develop clinical symptoms and the virus was cleared within 3 days. The
molecular components required for HuNoV infection, as well as the protective immunity
against HuNoV, cannot be identified using most animal models due to the difficulty in
genetically manipulating bovine, porcine and feline hosts when compared to mouse (Wobus
et al., 2006), emphasising the need to develop a tissue culture system.
To date, no cell culture systems have successfully propagated HuNoV, in spite of
several attempts using human and non-human cell culture (Duizer et al., 2004b; Papafragkou
et al., 2013). Only three studies by Straub and colleagues (Straub et al., 2011; Straub et al.,
2007; Straub et al., 2013) have had a limited success in cultivating GI and GII of human
NoVs in a 3-dimensional system using the human embryonic intestinal epithelial cell line
INT-407 and Caco-2 cells, but other workers have been unable to reproduce these findings
(Herbst-Kralovetz et al., 2013; Papafragkou et al., 2013; Takanashi et al., 2014). In other
studies, 19 cell culture models from 11 different animals were infected with a sample known
to contain HuNoV, but none of these cells were susceptible to HuNoV (Malik et al., 2005). In
addition, Duizer et al., 2004b used a variety of human and animal cell lines including, A549,
AGS, Caco-2, CCD-18, CRFK, CR-PEC, Detroit 551, Detroit 562, FRhK-4, HCT-8, HeLa,
HEC, HEp-2, Ht-29, HuTu-80, I-407, IEC-6, IEC-18, Kato-3, L20B, MA104, MDBK,
65
MDCK, RD, TMK, Vero and HEK293, as well as different laboratory methods, but all these
efforts were unsuccessful to propagate HuNoV in any of these cell lines (Duizer et al.,
2004b). Recently, a low level of HuNoV infection was detected in vitro using human B cells
in cell culture, however this required co-infection with enteric bacteria and therefore may not
be an ideal system (Jones et al., 2014). Therefore, most of the studies in recent years on
HuNoV are largely dependent on surrogate models (Vashist et al., 2009).
The animal caliciviruses are used as surrogates for the study of HuNoVs include
Sapovirus, porcine enteric calicivirus (PEC), feline calicivirus and Tulane virus (Rocha-
Pereira et al., 2014). The differences between these viruses and human noroviruses in regards
to genome organization, transmission and pathogenicity make them less than ideal for the
discovery of a HuNoV vaccine (Rocha-Pereira et al., 2014), but they provide an opportunity
to understand the pathogenicity and translation/replication mechanisms of HuNoV (Chaudhry
et al., 2006; Karst, 2010; Thorne & Goodfellow, 2014; Wobus et al., 2006).
FCV is frequently used as a model for the study of human noroviruses in regard to
environmental stability, survival, diagnostic and inactivation studies (Bae & Schwab, 2008;
Bozkurt et al., 2014; Buckow et al., 2008; Topping et al., 2009). This virus is also used as a
model to study human norovirus replication and translation due to the possibility of genetic
manipulation of the FCV genome (Chaudhry et al., 2006; Willcocks et al., 2004).
Discovery of murine norovirus in 2003 (Karst et al., 2003) and subsequent
propagation of this virus in tissue culture using cells of the hematopoietic lineage such as
macrophages (MΦ) and dendritic cells (DCs) (Wobus et al., 2004) provided an efficient cell
culture model for the study of many aspects of norovirus biology (Arias et al., 2012; Wobus
et al., 2006). At present, MNV is considered the best model for HuNoV due to its genetic
similarity, protein function similarity and ability to replicate in cell culture and in small
66
animal models, providing opportunities for specific genetic manipulation in the host (Karst et
al., 2003; Wobus et al., 2004; Wobus et al., 2006; Zheng et al., 2006). Therefore several
studies have used MNV as a model for understanding many aspects of HuNoVs life cycle
(Chaudhry et al., 2006; Thorne & Goodfellow, 2014; Vashist et al., 2009; Wobus et al.,
2006). MNV replication also occurs in many murine macrophage and dendritic cell-lines
including (RAW 264.7, IC21, J774A.1, P388D1, WBC264-9C and JAWSII) (Changotra et
al., 2009; Wobus et al., 2006). This raised the possibility that HuNoV might replicate in
similar cells of human origin and share a similar tropism to cells of the hematopoietic lineage
(Leung et al., 2010). However, Norwalk virus is unable to replicate in macrophage or
dendritic cells derived from the peripheral blood of susceptible humans suggesting that
human and murine norovirus tropisms differ (Lay et al., 2010). More recently, amino acid
changes in the genome of some MNV strains including MNV-3 and MNV.CR6 have been
associated with persistence and tropism (Arias et al., 2012; Nice et al., 2013). The MNV
model has provided new insight into how HuNoV interact with their host and has generated
questions regarding the relationship between noroviruses and cells of the hematopoietic
lineage and the role of innate immunity in the control of norovirus infection (Wobus et al.,
2006).
Recently, Tulane virus (TV) was isolated from stool samples of rhesus macaques, and
due to the ability to propagate this virus in tissue culture (Monkey kidney cell line LLC-
MK2), TV may provide another valuable model for understanding HuNoVs biology (Farkas
et al., 2008).
In addition to these models, the transfection of human hepatoma Huh-7 cells with
HuNoV RNA leads to virus replication but the newly formed viruses are unable to infect
other cells. This may be due to the absence of specific factors associated with viral entry
(Guix et al., 2007). A viral replicon system for NoV was generated by Chang and co-worker
67
(2006) and used to study a variety of potential antiviral molecules against HuNoV infection
(Bok et al., 2008; Chang & George, 2007; Chang et al., 2006). Recombinant virus like-
particles (rVLPs) of HuNoVs generated by expression of the major capsid protein (VP1),
with or without the minor capsid protein (VP2) (Lin et al., 2014; Vashist et al., 2009), serve
as a good model to study viral attachment (Harrington et al., 2002a) and provide opportunity
to understand the immunity against HuNoVs or develop antivirals (Harrington et al., 2002b;
Vashist et al., 2009).
1.11 Pathogenesis and immunity
Studies on NoVs pathogenicity and immunity have been hampered by the lack of a
reproducible culture system or small animal model for the propagation of human norovirus.
However, studies using FCV and MNV as models, as well as information from studies using
human volunteer have provided valuable information (Karst, 2010). Recently, animals such
as pigs and calves as well as non-human primates, infected with HuNoV, were used as
experimental animal models for understanding HuNoV biology (Bok et al., 2011; Cheetham
et al., 2006; Souza et al., 2008; Souza et al., 2007). Although viral RNA is detected in sera or
cerebrospinal fluid of infected volunteers, the upper part of intestinal tract (duodenum and
upper jejunum) remains the only target for NoVs infection (Ito et al., 2006; Takanashi et al.,
2009). Proximal intestinal biopsies have revealed specific histological changes in the
intestinal epithelium including broadening and blunting of the intestinal villi, shortening of
the microvilli, crypt-cell hyperplasia, and epithelial cell disarray and cytoplasmic
vacuolisation. In addition, infiltration of inflammatory cells (polymorphonuclear and
mononuclear cells) into the lamina propria is also observed (Troeger et al., 2009). In contrast,
no histological changes can be detected in the gastric fundus, antrum and colonic mucosa
68
(Levy et al., 1976). Gastric emptying is delayed during infection and this may be responsible
for nausea and vomiting (Meeroff et al., 1980). A transient malabsorption of fat, D-xylose
and lactose is also reported during infection and associated with the alteration of brush border
enzyme activity (alkaline phosphatase, sucrase and trehalase) in infected cells (Agus et al.,
1973). The susceptibility or resistance to infections depend primarily on the presence of
specific human HBGA receptors in the human gut (Lindesmith et al., 2008). Although more
than 80% of adults appear susceptible to HuNoVs infection, some adults display resistance to
infection due to a lack of HBGA receptors (Lindesmith et al., 2003). In addition, the
expression of HBGAs correlates with strain-specific susceptibility (Lindesmith et al., 2003).
HBGAs are a family of carbohydrates expressed in the mucosal surfaces of the respiratory,
genitourinary and digestive tracts, or linked to proteins/lipids on the red blood cell surface or
present as free oligosaccharides in biological fluids such as milk and saliva (Hutson et al.,
2004). They serve as receptors for NoVs and the binding to these receptors varies depending
on the strains (Lindesmith et al., 2008). Immunological studies reported that volunteers
display short-term immunity against NoV infections, varying from 4 to 16 weeks and
potentially as long as 2 years (Atmar et al., 2011; Johnson et al., 1990; Rocha-Pereira et al.,
2014). On average infected volunteers were repeatedly susceptible to infection with the same
strain or heterologous strains six months after previous exposure (Johnson et al., 1990;
Parrino et al., 1977). The viral dose is one of the many concerns with volunteer studies.
Although HuNoV has a very low infectious dose (less than 20 viral particles) as well as high
rate of shedding in stool (108-1010 RNA copies/gram), most studies challenged the volunteers
with doses several thousand-fold times higher than the minimum capable of causing human
illness (Teunis et al., 2008). More recently, Simmons et al., 2013 suggested that the duration
of immunity to infections lasts from about 4-8 years using six different mathematical models
69
to estimate immunity duration, depending on factors such as transmission process and natural
history of HuNoV (Simmons et al., 2013).
Both types of immunity, innate and acquired immunity have been reported to
contribute towards immunity against NoV infections (Parrino et al., 1977). The early
volunteers’ studies demonstrated no association between pre-existing serum antibodies and
the susceptibility to infection with Norwalk virus in adults (Atmar et al., 2011; Dolin et al.,
1971; Graham et al., 1994; Reeck et al., 2010). Volunteers with high levels of pre-existing
antibodies were more likely to have symptomatic illness compared to those with low levels of
antibodies (Johnson et al., 1990). In contrast, high level of antibodies may reflect short-term
immunity and recent exposure to HuNoV in children (Matsui & Greenberg, 2000). Using
MNV as a model in mice lacking functional type I interferon (IFN) signalling pathways,
studies demonstrated the requirement of type I IFN immune protection against MNV
infection (Changotra et al., 2009; Karst et al., 2003; Mumphrey et al., 2007). In addition,
(Jung et al., 2012) showed that the oral administration of type 1 IFN to gnotobiotic pigs
infected with MNV led to decrease viral shedding. IFN acts via activated STATs causing a
cascade of events leading to the induction of antiviral proteins such as RNA-dependent
protein kinase (PKR) and RNase L (Samuel, 2001). A study using a Norwalk replicon system
demonstrated that norovirus replication is inhibited by type I IFN and type II IFNγ (Chang
& George, 2007; Chang et al., 2006). The role of T-cells in this process is not completely
understood, but studies have reported that HuNoV infection or vaccination using virus-like
particles (VLPs) elicit CD4+ Th1 response leading to an increase of IFNγ and IL-2 cytokine
secretion (Lindesmith et al., 2005; Lindesmith et al., 2008). Recently, Troeger et al., 2009
observed an increase in the number of intraepithelial cytotoxic T cells in the duodenum
during the first six days of infection (Troeger et al., 2009). Moreover, this study also
correlated the apoptosis of human enterocytes with HuNoVs infections. The increase in Th1
70
as well as Th2 cytokines and interferons was also reported in serum and intestinal contents of
gnotobiotic pigs and calves infected with human GII.4 strain of HuNoVs (Souza et al., 2008;
Souza et al., 2007).
Despite the complex nature of protective immunity to NoVs infection, due to the
antigenic diversity of HuNoVs, and also the lack of an efficient culture model, recent
attempts to produce a vaccine against HuNoVs using virus-like particles (VLP) have shown
promise (Herbst-Kralovetz et al., 2010; Karst, 2010; Karst et al., 2014a; Rocha-Pereira et al.,
2014; Rohayem et al., 2010; Thorne & Goodfellow, 2014; Tome-Amat et al., 2014).
1.12 Viral structure, genome organisation and function of the viral proteins
Caliciviruses are small non-enveloped spherical viruses between 28-38 nm in
diameter containing a single-stranded, positive-sense, polyadenylated RNA genome, whose
organisation varies among the calicivirus genera (Figure 1.9). While Sapovirus, Nebovirus
and Lagovirus genera have a genome consisting of two open reading frames (ORF1 and
ORF2), the genomes of Vesivirus and Norovirus are mainly organised into three ORFs (Jiang
et al., 1993; Rohayem et al., 2010). The non-structural proteins in all caliciviruses are
encoded by ORF1, but the expression of the structural proteins, the major capsid protein
(VP1) and minor capsid protein (VP2) differ among the genera. In Vesivirus and Norovirus,
VP1 and VP2 are encoded by ORF2 and ORF3, respectively; whereas in Sapovirus,
Lagovirus and Nebovirus genera, VP1 and VP2 are encoded by ORF1 and ORF2,
respectively (Rohayem et al., 2010). A common feature of all caliciviruses is the lack of a
methylated cap (m7GpppN) at the 5´ terminus of the viral RNA genome. Instead, a 10-15 kDa
protein named as VPg (Virion Protein genome-linked) is attached to the 5´end and serves as a
cap substitute for initiation of viral protein translation (Burroughs & Brown, 1978; Herbert et
71
al., 1996; Karst et al., 2014a; Meyers et al., 1991). The calicivirus genome is polyadenylated
at its 3´end, terminating with a poly (A) tail of 110 nucleotides (nt) (Burroughs & Brown,
1978; Farkas et al., 2008; Jiang et al., 1993; Xi et al., 1990) (Figure 2). The mature
calicivirus virions are composed of three proteins including VP1, VP2 and VPg (Sosnovtsev
et al., 2000). While VPg is found in mature virions it only functionally serves as a non-
structural protein during replication (Green, 2013).
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Figure 1.9 Organization of representative caliciviruses genome. The caliciviruses genome is
covalently linked to VPg (NS5) at the 5’ and with a poly(A) tail and is organised into 2-4 ORFs as
indicated (A, B, C, D). ORF1 is translated as a polyprotein, cleaved by the protease NS6 at
different cleavage sites to produce the non-structural proteins that are essential for viral replication
(A, B, C, and D) and VP1 (D). VP2 is encoded by ORF3 in (A, B and C) or by ORF2 in (D).
ORF2 and ORF3 are translated from a subgenomic RNA in all caliciviruses. In addition MNV (B)
has an additional alternative fourth ORF. ORF4 overlaps with ORF2 and is also translated
primarily from the subgenomic RNA into the virulence factor 1 (VF1) protein.
73
Noroviruses genome range from 7.3-7.7 kb (kilobases) in size organised into three
ORFs (Thorne & Goodfellow, 2014; Zheng et al., 2006). However, some strains of MNV
contain a fourth small ORF, ORF4, which overlaps ORF2 and is encoded by a sub-genomic
RNA (sgRNA). ORF4 encodes a 214 amino acid protein called virulence factor 1 (VF1),
involved in the regulation of the host innate immunity response and apoptosis (McFadden et
al., 2011; Thackray et al., 2007). ORF1, the largest ORF (5 kb) encodes the large poly-
protein precursor (~ 200 kDa) which is self-cleaved during translation to yield six/seven non-
structural viral proteins (NS1-7) by the viral proteinase (Pro; NS6) in the following order: the
N-terminal protein (p48; NS1-2 in MNV;37-48 kDa), the 2C-like nucleoside triphosphatase,
NTPase (NS3; ~40kDa), picornavirus 3A-like protein (p22; NS4 in MNV; 20–30 kDa), VPg
(NS5), the viral 3C-like protease 3CLpro (Pro; NS6 in MNV;19 kDa) and RNA-dependent
RNA polymerase (RdRp; NS7 in MNV; ~57 kDa) (Belliot et al., 2003; Clarke & Lambden,
2000; Hardy, 2005; Seah et al., 2003; Sosnovtsev et al., 2006).
ORF2 (1.8 kb) and ORF3 (0.6 kb) encode the structural proteins, the major capsid
protein (VP1; 57 kDa) and minor capsid protein (VP2; 22 kDa) respectively (Donaldson et
al., 2008; Glass et al., 2000a; Green et al., 2000; Jiang et al., 1992). Both of them are
encoded by a sub-genomic RNA (sgRNA; ~ 2.3 kb) (Katayama et al., 2006; Neill &
Mengeling, 1988; Wobus et al., 2004). VP2 is a small protein found in only few numbers of
copies per virion with yet unknown function (Thackray et al., 2007), but some data suggest
that VP2 may play a role in increasing the stability of VP1 by protecting it from protease
degradation (Bertolotti-Ciarlet et al., 2003), or in encapsidation of the viral genome (Karst,
2010; Vongpunsawad et al., 2013). VP2 is required for the formation of stable virus-like
particles (VLPs) via its interaction with VP1 (Bertolotti-Ciarlet et al., 2002; Di Martino &
Marsilio, 2010). VP1 protein, the main capsid protein, is composed of three domains linked
74
by a flexible hinge including the N domain, the internal part of the capsid, the S domain
forming the icosahedral shell and a third variable P domain, which contains two sub-domains
P1 and P2 forming the protrusions (Allen et al., 2008; Chen et al., 2006; Prasad et al., 1999;
Tan et al., 2008) (Figure 1.10). The P2 domain is the most hypervariable region of the capsid
which is located at the outermost capsid surface and is responsible for cellular binding
(especially to HBGAs) and immune recognition (Allen et al., 2008; Cao et al., 2007;
Chakravarty et al., 2005; Chen et al., 2006; Glass et al., 2000a; Hardy, 2005; Prasad et al.,
1999; Tan et al., 2003) (Figure 1.10B and C). Furthermore, the receptor binding specificity of
HuNoV seems to be associated with the P2 sub-domain (Prasad et al., 1999; Tan et al., 2008;
Tan et al., 2003). Therefore, this subdomain represents one of the best targets for determining
the strains that correlate with outbreaks (Lindesmith et al., 2013; Sukhrie et al., 2013; Vega
et al., 2011; Xerry et al., 2009). The 3-dimensional structure of the viral capsid determined
using X-ray crystallography showed that the virion is formed of 180 molecules of the VP1
distributed into 90 dimeric capsomeres associated with a few molecules of VP2 (usually one
or two copies) (Prasad et al., 1999; Vongpunsawad et al., 2013). The capsid protein is also
critical for viral replication through a novel interaction with the RdRp (Morillo & Timenetsky
Mdo, 2011; Subba-Reddy et al., 2012).
The function of some non-structural proteins remains unknown. Some studies have
suggested that N-terminal proteins may play a role in membrane rearrangement and
intracellular protein trafficking (Ettayebi & Hardy, 2003; Fernandez-Vega et al., 2004). NS4
and the NTPase have been shown to have functional similarity to the picornaviruses 2C and
3A protease, respectively (Hyde et al., 2009; Sosnovtsev et al., 2006). NS1-2, NS3 and NS4
proteins contribute to processes essential for MNV replication such as membrane induction or
cellular manipulation (Hyde et al., 2009). The NS6 protease shares structural similarity with
the cellular chymotrypsin-like serine proteases (Blakeney et al., 2003), and share functional
75
and structural properties with picornavirus 3C proteases (Leen et al., 2012). The NoVs
proteases play an essential role in the maturation of the ORF1 polyprotein precursor into
mature non-structural proteins via cleavage of the polyprotein at different cleavage sites (QG,
EG and EA), producing six active proteins (Liu et al., 1996; Sosnovtsev et al., 2006;
Takahashi et al., 2013a; Takahashi et al., 2013b). In addition, NS6 also plays a role in the
inhibition of cellular translation via cleavage of both eukaryotic initiation factor 4G and poly
(A) binding protein (critical for initiation of translation) (Kuyumcu-Martinez et al., 2004;
Zeitler et al., 2006). In FCV, p76 acts as a protease and polymerase. The first half of p76 acts
as a trypsin-like serine protease and is related to picornavirus 3C-protease, therefore it is
designated 3C-like protease, while the second half of p76 is related to picornaviruses 3D-
polymerase and acts as an RNA-dependent-RNA polymerase (RdRp). In contrast, in MNV
the p76 is separated into two proteins including NS6 which acts as protease, and NS7 that
acts as a polymerase (Sosnovtsev et al., 2006). The crystal structure of RdRp from GII
noroviruses displayed general similarity to the structural domains of other positive strand
RNA viruses, especially with RHDV (Hardy, 2005). VPg protein (NS5) was discovered in
the late 1970s and is a small 10-15 KDa protein linked to the 5´ end of genomic and sub-
genomic RNAs in all members of Caliciviridae, and similar proteins are found in other
positive sense RNA viruses such as the Picornaviridae, Potyviridae, Luteoviridae,
Comoviridae and Secoviridae, although with different functions (Burroughs & Brown, 1978;
Clarke & Lambden, 2000; Dunham et al., 1998; Herbert et al., 1996; Kneller et al., 2006;
Kozak, 1991; Sadowy et al., 2001). For instance the VPg of picornaviruses is not involved in
translation initiation (Fitzgerald & Semler, 2009; Goodfellow, 2011), but plays a role in
replication whereas the VPg of Potyvirus (Lellis et al., 2002; Leonard et al., 2000) and
caliciviruses (Daughenbaugh et al., 2006; Herbert et al., 1997) is important for protein
synthesis. (Daughenbaugh et al., 2003) showed a direct interaction between VPg and
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eukaryotic initiation factor 3 (eIF3) of HuNoV suggesting the involvement of VPg in
translation initiation. Further evidence for the role of VPg in translation initiation came from
Goodfellow et al., 2005 who reported that FCV VPg interacts with eukaryotic initiation
factor 4E (elF4E) in infected cells and that this interaction is necessary for translation of FCV
in vitro (Goodfellow et al., 2005). This interaction between VPg and eIF4E has also been
reported in MNV-1 (Daughenbaugh et al., 2006) but the role of this interaction VPg-eIF4E
in protein synthesis is less clear (Chaudhry et al., 2006; Goodfellow, 2011). The infectivity of
caliciviruses is reduced by removal of the VPg protein from 5′end of the mRNA (Chaudhry et
al., 2006; Guix et al., 2007). In addition, VPg may play a possible role in transporting the
genome to the site of negative strand synthesis (Donaldson et al., 2008).
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Figure 1.10 The structure of norovirus major capsid protein VP1. (A) The capsid
protein is encoded by open reading frame 2 (ORF2). VP1 consists of the shell (S) and
protruding domains (P), linked by a flexible hinge, and P is divided into two subdomains,
including P1 (blue) and P2 (red). (B, C, D) P2 is located at the capsid surface and is
responsible for cellular binding and immune recognition.
78
1.13 Norovirus life cycle
The exact mechanisms for HuNoV translation and replication are yet to be determined
and most knowledge originates from studies of animal surrogates (Donaldson et al., 2008;
Karst et al., 2014a; Thorne & Goodfellow, 2014; Vashist et al., 2009). Norovirus replication
occurs in the cytoplasm of the host cell, possibly enterocytes (Cheetham et al., 2006; Karst,
2010). The replication cycle of noroviruses consists of several steps (Figure 1.11) beginning
with the attachment of the VP1 P2 domain to specific cellular receptors in the surface of host
cells which in the case of HuNoVs, are the HBGAs (Abrantes et al., 2012b; Hutson et al.,
2003; Marionneau et al., 2002; Tan & Jiang, 2005). Caliciviruses such as rhesus enteric
calicivirus (ReCV), RHDV and canine noroviruses also interact with their host cells via
HBGA receptors (Caddy et al., 2014; Farkas et al., 2010; Ruvoen-Clouet et al., 2000; Tian et
al., 2007). In contrast, the interaction of other members of caliciviruses with host cells
occurs via different cellular receptors. For instance, FCV attaches to the host cell membrane
via junction adhesion molecule-1 (JAM-1) (Makino et al., 2006), whereas MNV uses a
terminal sialic acid on ganglioside GD1a of a mouse macrophage, glycolipids and
glycoproteins as a receptor depending on the MNV strains (Taube et al., 2012; Taube et al.,
2009).
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Figure 1.11 Proposed replication mechanism of Human Norovirus (HuNoV). The replication
cycle can be divided into the following stages: (1) attachment of virus via P2 to the cellular
receptor, (2) entry and (3) uncoating of the viral genome, (4, 5, 6 & 7) translation of genomic RNA
into a polyprotein precursor through VPg-dependent translation (4 & 5), followed by post-
translational processing mediated by protease (Pro) to produce six non-structural proteins (6) which
are released and accumulate in replication complex (RC) where replication occurs (7), (8, 9 & 10)
RNA replication is mediated by Pol and other non-structural proteins in RC to generate anti-
genomic RNA (negative-strand RNA) (8), which used as a template for the synthesis genomic RNA
as well as sub-genomic RNA translated as structural proteins (VP1 & VP2) (9 & 10), (11) assembly
and maturation of the structural proteins as well as packaging of the VPg-linked RNA genome
occurs, followed by release and exit of the mature virion from the cell (12).
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Many studies have reported the role of specific HBGAs in attachment of different
genotypes of NoV and recombinant virus-like particles (VLPs) to epithelial host cells,
especially Caco-2 cells (Harrington et al., 2004; Hutson et al., 2003; Marionneau et al., 2002;
White et al., 1996). The viral attachment is associated with H type 1 antigen which produces
the FUT2 enzyme (Marionneau et al., 2002). Herbst-Kralovetz et al., 2013 have reported the
lack of expression of H type 1 and 2 or Lewis b antigens on the differentiation of Int-407
cells in a 3-D cell culture system, which may explain why the human macrophage and
dendritic cells were unable to replicate HuNoVs (Papafragkou et al., 2013). Moreover, the
CEP-like morphological change of the 3-D aggregates reported by Straub et al., 2007 may be
due to the presence of lipopolysaccharides or other stool-contaminating toxins (Herbst-
Kralovetz et al., 2013). Epidemiological studies have provided strong evidence for the
requirement of NoV infection for HBGAs as receptors or co-factors (Hennessy et al., 2003;
Hutson et al., 2002; Lindesmith et al., 2003). There are two binding profile groups of NoVs,
with strains that bind A/B and/or H antigens and strains binding Lewis and/or H antigens. In
addition, another group of strains does not bind to HBGAs raising the possibility that other
receptors can be used by NoVs for infection (Donaldson et al., 2008; Murakami et al., 2013).
More recently, Murakami et al., 2013 demonstrated that the binding of NoV-like particles to
human intestinal epithelial cells line (Caco-2) occurs independently of HBGAs but depends
on the state of cell differentiation. This suggested that VLPs utilize different cell receptors or
co-receptors during binding and internalization into cells (Murakami et al., 2013).
Following attachment, the viruses enter the host cells through an unknown
mechanism. In the case of MNV, internalisation occurs by a non-clathrin, non-caveolin
mediated endocytic pathway which depends on dynamin and cholesterol (Gerondopoulos et
al., 2010; Perry & Wobus, 2010). The entry of FCV (F9 strain) occurs via clathrin mediated
endocytosis in a pH-dependent manner (Kreutz & Seal, 1995; Stuart & Brown, 2006). After
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genome uncoating, and release of the viral genome into the cytoplasm of the host cell, protein
synthesis begins with recognition of the VPg-linked RNA genome by the translation
machinery of the host cells (Rohayem et al., 2010; Thorne & Goodfellow, 2014). Caliviruses
use a novel mechanism for the initiation of protein synthesis: a VPg-dependent translation
initiation involving the interaction of eukaryotic initiation factors (eIFs) with VPg protein,
acting as a cap substitute (Chaudhry et al., 2006; Daughenbaugh et al., 2003; Goodfellow et
al., 2005; Herbert et al., 1997) (Figure 1.12). These interactions lead to recruitment of the
ribosome and host translation factors to the viral RNA and thereby initiation of translation
(Chaudhry et al., 2006; Daughenbaugh et al., 2003; Goodfellow et al., 2005).
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Figure 1.12 Schematic diagram of cap and VPg-dependent translation initiation. (A) The cap-
dependent translation mechanism is dependent of the recognition of 5’ m7Gppp(N) cap structure by the
eIF4F complex consistent of the cap binding protein eIF4E, the scaffolding protein eIF4G, the RNA helicase
eIF4A and its co-factor eIF4B. Subsequently eIF4G mediates the recruitment of the 40S ribosomal subunit
and other eIFs via an interaction with eIF3, this leads to the scanning mechanism that then locates the AUG
start codon. (B) In contrast, during VPg-dependent translation, VPg acts as a cap substitute and interacts
with eIF4E, eIF4G and eIF4A to tether ribosomes to the 5’ end of the viral RNA. During FCV infection, the
interaction of VPg with eIF4E and eIF4G is required for translation while only the interaction with eIF4G is
required during MNV infection.
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Translation of ORF1 of the viral genome produces a large single polyprotein that
subsequently undergoes auto-proteolytic processing mediated by the viral protease (3CLPro or
NS6) to release mature NS proteins and their precursors (Sosnovtsev et al., 2006). Five
3CLPro cleavage sites have been reported in noroviruses and in the case of MNV-1 NS
proteins these sites include 341E|G342, 705Q|N706, 870E|G871, 994E|A995, 1177Q|G1178 (Sosnovtsev
et al., 2006). These NS proteins along with host proteins and other viral RNA molecules
assemble in intracellular membranous structures called replication complexes (RC) where
genome replication occurs (Green et al., 2002; Hyde et al., 2009; Wobus et al., 2004). In the
case of MNV, the NS proteins p48 (NS1-2) and p22 (NS4) are required to recruit host
membranes (endoplasmic reticulum (ER), Golgi complex and endosome) and induce RC
formation, providing a platform for replication (Hyde & Mackenzie, 2010; Hyde et al., 2009;
Sharp et al., 2010) via the modulation of the host cell secretory pathway (Denison et al.,
2008). In addition, HuNoVs p48 and p22 proteins (equivalent to the MNV NS1-2 and NS4
respectively) induce disassembly of the Golgi complex and inhibit protein secretion, therefore
these proteins may have a possible role in driving cellular membrane rearrangement (Ettayebi
& Hardy, 2003; Fernandez-Vega et al., 2004; Sharp et al., 2010). Within the RC, the genomic
RNA serves as a template for anti-genomic RNA (negative sense) production via a
transcriptional process mediated by the RdRp which uridylates the VPg in the presence of
polyadenylated genomic RNA leading to the production of the negative-sense strand
(Rohayem et al., 2010). This anti-genomic RNA is then used as a template for the synthesis
of new genomic RNA, which is either translated or packaged into new virions, and sub-
genomic RNA that is translated into structural proteins (VP1 and VP2). The neo-synthesized
genomes are packaged into the capsid to assemble a mature virion which is finally released
from the cell via yet unknown mechanisms (Rohayem et al., 2010).
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The exact mechanism of sgRNA production is still unclear, but in the case of MNV
two possible mechanisms have been suggested, both dependent on RdRp. The first is via
internal initiation involving the binding of the viral RdRp to the sgRNA promoter located
downstream of the sgRNA start site in the negative strand followed by synthesis of a VPg-
linked positive strand sgRNA (Miller et al., 1985). In the second model, the RdRp binds to
the 3′ end of the positive strand genomic RNA and initiates negative-strand synthesis, and
then RdRp terminates at a specific sequence located at the 3′ end of the negative strand
sgRNA leading to production of negative strand sgRNA, which then acts as a template to
produce positive strand sgRNA (Sit et al., 1998). Other studies demonstrated the role of
RdRp of HuNoV and MNV in this process by recognising and binding to a sgRNA promoter
(Lin et al., 2015). Using this sub-genomic RNA as template, the translation of VP2 happens
through termination/reinitiation process, which depends on an upstream sequence named
termination upstream ribosomal binding site (TURBS), a 40–70 nt termination upstream
ribosomal binding site (Luttermann and Meyers, 2014).
1.14 The function of RNA-protein interaction during noroviruses infections
In addition to interactions with the host translation machinery (Figure 1.12), other
cellular proteins and conserved viral RNA structures or sequences also play an important role
in many aspects of viral life cycle (Liu et al., 2009; Lopez-Manriquez et al., 2013; Simmonds
et al., 2008; Thorne & Goodfellow, 2014; Vashist et al., 2012). Many studies have reported
direct and indirect protein-protein or RNA-protein associations between virus and their host,
some of which are required for the norovirus life cycle (Chaudhry et al., 2006; Chung et al.,
2014; Goodfellow et al., 2005; Gutierrez-Escolano et al., 2000; Karakasiliotis et al., 2010;
Lopez-Manriquez et al., 2013; Vashist et al., 2012).
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VPg forms several possible interactions with elements of the translation initiation
complex, but not all these interaction required for viral translation or replication (Chaudhry et
al., 2006; Vashist et al., 2012). These studies reported a direct or indirect association between
calicivirus VPg and specific eukaryotic initiation factors such as eIF4F components (4A, 4E
and 4G) as well as eIF3 (Chaudhry et al., 2006; Daughenbaugh et al., 2003; Goodfellow et
al., 2005). For instance, in vitro studies using reticulocyte lysate have demonstrated a direct
interaction between eIF4E (cap-binding protein) and VPg during MNV infection but this
interaction is not essential for MNV translation (Chaudhry et al., 2006; Goodfellow et al.,
2005). In contrast, the same interaction is critical in the case of FCV, as translation is
inhibited in vitro either by depletion of eIF4E or inhibition of the eIF4E-eIF4G1 interaction
(Chaudhry et al., 2006). However, the eIF4G-VPg association mediated by the C-terminal
fragment of eIF4G during MNV infection is necessary for translation and efficient viral
replication (Chung et al., 2014). Furthermore, a direct interaction between VPg and eIF3 has
been reported, but the role of this interaction is unknown (Daughenbaugh et al., 2003).
Additional indirect associations between viral VPg and other cellular proteins such as
LARP1, ABCE1, DDX9, IGFBP1, eEF1A1 and different ribosomal proteins were also
reported but the role of these interactions remain unclear (Chung et al., 2014). In vitro studies
also demonstrated that the RNA helicase eIF4A is required for MNV and FCV translation for
unwinding the secondary structure in the 5’ end of RNA, and inhibiting this initiation factor
either by using a dominant-negative form of eIF4A or a small molecule inhibitor such as
hippuristanol impairs both MNV and FCV translation (Chaudhry et al., 2006; Simmonds et
al., 2008). More recently, Royall et al., 2015 demonstrated that phosphorylation of eIF4E,
occurring via activation of the MAPK pathway is required for MNV1 replication (Royall et
al., 2015).
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Vashist et al., 2012 identified approximately 150 different proteins that interact with
the 5’ and 3’ extremities of the MNV-1 genome using rabbit reticulocyte lysate, but the role
of most of these interactions remains elusive requiring further additional studies (Vashist et
al., 2012). These interactions between the 5’ and 3’ends of the viral genome with host
proteins include Lupus autoantigen (La), polypyrimidine tract binding protein (PTB) and poly
(A) binding protein (PABP) (Vashist et al., 2012) which play an important role in MNV
replication (Gutierrez-Escolano et al., 2000; Gutierrez-Escolano et al., 2003; Kuyumcu-
Martinez et al., 2004). These studies also reported the interaction of La and polypyrimidine
tract binding protein (PTB) with both 5’ and 3’extremities of the Norwalk virus genome
(Gutierrez-Escolano et al., 2000; Gutierrez-Escolano et al., 2003). La protein, which
regulates the translation of other RNA viruses such as Hepatitis C Virus (HCV) (Kumar et
al., 2013; Ray & Das, 2011), was also reported to interact with HuNoV genome (Fitzgerald
& Semler, 2009). In the case of FCV, the interaction of PTB with the 5’ end of genomic and
sub-genomic RNA, as well as its association with RC, has been demonstrated to be essential
for viral replication (Karakasiliotis et al., 2010). In contrast, the cellular proteins La, PTB and
DDX3 have a negative impact on MNV replication in vitro (Vashist et al., 2012). Because of
the role of PTB in promoting MNV translation, but reducing translation of related viruses
such as FCV, the possibility has been raised that it plays a role in switching from translation
to replication (Karakasiliotis et al., 2010; Vashist et al., 2012).
The binding of PABP to poly (A) tails at the 3´ end of mRNAs plays a key role in
mRNA translation and stability by facilitating mRNA circularization via its binding to
eIF4G (Gorgoni & Gray, 2004). This closed loop is believed to stimulate translation (Imataka
et al., 1998; Sonenberg & Hinnebusch, 2009), and is important for promoting ribosome
cycling and stimulating 60S subunit joining (Kahvejian et al., 2005). However later in
infection, the calicivirus 3CLPro inhibits cellular translation by cleaving PABP (Kuyumcu-
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Martinez et al., 2004; Willcocks et al., 2004). Recently, Lopez-Manriquez et al., 2013
identified an important role for poly(C) binding protein (PCBP) and hnRNPA1 which bind to
5’ end of MNV-1 genome promoting genome circulation and viral replication (Lopez-
Manriquez et al., 2013). These studies also demonstrated the association of the 5’ end with
PCBP-2 and hnRNPA1 or PABP with the 3’ end. The role of these interactions remains
elusive but comparison with other positive-strand RNA viruses such as poliovirus and dengue
virus suggests a possible role in the circulation of the viral genome (Vashist et al., 2012).
88
1.15 Aims and objectives of the research
It is clear from the previous section that calicivirus replication is dependent on a
complex network of interactions between the viral RNA and the host translation machinery to
mediate viral protein synthesis and to regulate translation in the host. In addition, many
studies have demonstrated that viral infections are a major stress for the host cell leading to
the regulation of mRNA translation at the level of cytoplasmic RNA granules.
Therefore the following hypothesis has been formulated: calicivirus replication
regulate the host response to infection by affecting the formation or assembly of stress
granules and P-bodies.
To address this hypothesis the following objectives have been designed:
- Using cell lines supporting FCV and MNV-1 replication to study the assembly of stress
granules and P-bodies using appropriate immunofluorescence microscopy markers.
- To dissect the effect of FCV and MNV-1 infection on the assembly of stress granules.
- To dissect the effect of FCV and MNV-1 infection on the assembly of P-bodies.
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Chapter 2
General Materials and Methods
90
2.1 Maintenance of cells:
2.1.1 Crandall Rees Feline Kidney (CRFK) Cells
CRFK cells were obtained from the European Collection of Cell Cultures (ECACC)
and grown in Eagle’s Minimum Essential Medium (MEM) with Earle’s salts, supplemented
with 10% (v/v) heat inactivated foetal bovine serum (FBS), penicillin/streptomycin 1% (v/v)
and non-essential amino acids 1% (v/v) (all from GibcoBRL, Life Technologies). Cells were
maintained in a 75-cm2 tissue culture flask at 37ºC with 5% CO2 (Incubator Galaxy S; Wolf
Laboratories), and passaged as follows: the medium was removed and the cells were rinsed
with 5 ml 1:10 trypsin/versene (T/V) (all from GibcoBRL life technologies). The T/V
solution was removed and cells were detached by addition of 5 ml trypsin/versene and the
flask was incubated at 37oC with 5% CO2 for 3 min. The suspension of cells and T/V was
transferred to a Universal tube containing 5 ml of 10% FBS-Eagle’s MEM and the cells were
centrifuged at 2500 rpm (Centurion) for 3.5 min. The medium was removed and the pellet
was resuspended in 6-ml 10% FBS-Eagle’s MEM. Approximately 1-1.5 ml of cell suspension
were placed in a new 75-cm2 flask containing 14 ml 10% FBS-Eagle’s MEM. The CRFK
cells were used for all experiments were obtained between passages numbers 6 -16.
2.1.2 Mouse Leukemic monocyte-macrophage Cells (RAW 264.7)
RAW 246.7 (ECACC) cells were grown in Dulbecco’s Minimum Essential Medium
(DMEM) with Earle’s salts, supplemented with 10% (v/v) FBS, penicillin/streptomycin 1%
(v/v) (all from GibcoBRL life technologies). Cells were maintained at 37ºC with 5% CO2
(Incubator Galaxy S; Wolf Laboratories), and passaged as follows: the medium was removed
and the cells were resuspended in 8 ml 10% FBS-Eagle’s DMEM in the presence of sterile
glass beads to detach the cells from the flask. Approximately 1-1.5 ml of cell suspension
91
were placed in a 75-cm2 flask containing 14 ml 10% FBS-Eagle’s DMEM. The cells were
used for all experiments were obtained between passages numbers 6 -16.
2.1.3 Feline embryonic airway (FEA) cells.
FEA cells were kindly provided from Dr Alan Radford (University of Liverpool, UK)
and grown in Eagle’s MEM with Earle’s salts, supplemented with 10% heat inactivated FBS,
1% penicillin/streptomycin and 1% non-essential amino acids. Preparation of confluent
monolayers was achieved as CRFK maintenance protocol (section 2.1.1).
2.1.4 Murine microglial cells (BV-2)
Murine microglial cells (BV2) were kindly provided from Prof Ian Goodfellow
(Cambridge University, UK). The cells were cultured in DMEM supplemented with 10%
(v/v) heat inactivated FBS, 1% penicillin/streptomycin, 1% L-glutamine and 1% sodium
bicarbonate (Life Technologies). The cells were incubated at 37ºC with carbon dioxide (5%)
in a Galaxy S incubator (Wolf Laboratories). Cells reaching a confluence of 80% were
scraped from the 75-cm2 flask substrate with sterile glass beads and centrifuged at 2500 rpm
(Centurion) for 3.5 min. The supernatant was removed and the pellet was resuspended in 8 ml
of growth medium after which 1 ml of the cell suspension was transferred into a 75-cm2
vented flask containing 14 ml of growth medium and grown at 37ºC with 5% CO2 (Incubator
Galaxy S, Wolf Laboratories).
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2.1.5 Human embryonic kidney 293 (HEK293T) cells (293 cells)
293 cells were grown in DMEM containing 1000 mg/L glucose (low glucose DMEM)
supplemented with 10% (v/v) foetal calf serum (FCS), 1% penicillin and streptomycin (Life
Technologies), 2% glutamine and 1% non-essential amino acids. The cells were incubated at
37ºC with 5% CO2 and then passaged when the confluence reached 80-90%. Preparation of
confluent monolayers was achieved as follows; the media was removed and the cells washed
with 3 ml 1% trypsin/versene (T/V) to remove remaining serum. Then, cells were detached
from the flask with 5 ml of fresh T/V. The detached cells were transferred to a 25 ml
universal tube containing 5 ml of fresh growth medium and centrifuged at 2500 rpm
(Centurion) for 3.5 min. The supernatant was removed and the cell pellet resuspended in 6 ml
of growth medium. 2 ml of the cell suspension was transferred to a 75-cm2 flask containing
16 ml fresh growth medium and incubated at 37ºC with 5% CO2 (Incubator Galaxy S, Wolf
Laboratories). Cell differentiation using poly-lysine (Sigma) treatment was applied for
culturing the cells on 13-mm coverslips as fellow; coverslips in 24-wells plate were treated
with poly-lysine solution, rocked and left at RT for 5 min. Poly-lysine were then removed
and coverslips were rinsed with sterile MQ water for several time before leaving the
coverslips to dry in a sterile hood for 2h and cells were then grown on coverslips.
2.1.6 Murine macrophage J774 cells
J774 cells (from the ATCC) were maintained in DMEM medium with 10% FBS at
37°C with 5% CO2. For sub-culturing, cells with confluence of 80-90% were scraped from
the 75-cm2 flask substrate with a cells scraper (VWR), spin down at 2500 rpm (Centurion) for
3.5 min and resuspended in 6-ml of fresh growth medium. 1ml of cell suspension was added
to 75-cm2 flask containing 14 ml of fresh growth medium.
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2.1.7 THP-1 cells
THP-1 cells (from the ATCC) were grown in RPMI 1640 medium (from ATTC)
supplemented with 2 mM L-glutamine, 0.05 mM beta-mercaptoethanol and 10% FBS. When
the cells were approximately 80% confluent, they were harvested, centrifuged at 1500 rpm
(Centurion) for 5 min and then suspended in 5-ml of fresh growth medium. I-ml of cell
suspension was added to 19-ml of growth medium and incubated at 37ºC and 5% CO2. Cell
were differentiated using PMA treatment prior immunofluorescence studies as fellow; Stock
of 5 mg/ml of PMA was prepared in DMSO and 4 µl of this stock was added to T-75 flask
containing 2-4 X 105 cells in 20 ml of media (5mg/ml). Flasks were then incubated overnight
in 37C and 5% CO2. For 3 days, following incubation, differentiated cells were washed with
PBS (3X). The PBS were then replaced by growth media.
2.2 Freezing and Resuscitation of CRFK and RAW264.7 cells
Freezing medium, 10 ml of appropriate growth medium for CRFK or RAW264.7
cells, 1 ml FBS and 1 ml dimethylsulphoxide (DMSO) was prepared for freezing cells as
follows: cells were harvested from one or more 75-cm2 flasks and subjected to centrifugation
at 2500 rpm (Centurion) for 3-4 min. The supernatant was removed and the pellet
resuspended in 2 ml freezing medium per flask and stored as 1ml aliquots in cryovials. The
cryovials were then placed overnight at -800C to freeze slowly. Next day, the cells were
transferred to liquid nitrogen storage.
For resuscitation, cells were thawed either at room temperature or rapidly in a 37ºC
water bath. The cells were then transferred to a universal tube containing 5 ml of the
appropriate warm (37ºC) medium and subjected to centrifugation at 2500 rpm (Centurion) for
3-4 min. The supernatant was discarded and the pellet resuspended in 7 ml of the appropriate
94
medium and transferred to a small flask (25-cm2). Following overnight incubation at 37ºC
with 5% CO2 (Incubator Galaxy S; Wolf Laboratories), the medium was exchanged for new
growth medium to remove any residual DMSO and the cells were incubated until confluent.
They were then passaged as normal (see section 2.1.1 and 2.1.2).
2.3 Preparation of viruses stock
2.3.1 Preparation of Feline Calicivirus (FCV) stock
A 175-cm2 flask of cells (80% confluent) was infected with 400µl of FCV Urbana
(obtained from Prof Ian Goodfellow, Cambridge University, UK) in 4 ml 2% (v/v) FBS-
Eagle’s MEM and rocked for 1 hour at room temperature (RT). After rocking, the medium
was removed and replaced with 8 ml of 10% (v/v) FBS-Eagle’s MEM and the cells incubated
at 37ºC for a further 5 hours before leaving the cells to freeze overnight at -80oC. The cells
were then thawed and scraped off and placed in a 20 ml universal, pelleted at 2500 rpm
(Centurion) for 2.5 min and the supernatant filtered through a 0.2 µm Millex filter, divided
into aliquots and stored at -80ºC. The virus titre was estimated by determination of the 50%
tissue culture infectious dose (TCID50) (Section 2.4).
2.3.2 Production of Murine Norovirus (MNV) stock
A confluent (approximately 80%) 75-cm2 flask of RAW264.7 cells was infected with
1-ml of MNV-1 (obtained from Prof H. Virgin, Washington University, St Louis, USA) seed
stock in 4 ml 2% (v/v) FBS-Eagle’s DMEM and rocked for 1h at RT. Another 8 ml of 10%
FCS- Eagle’s DMEM were added to the flask and incubated for 17 h until the cytopathic
effect was evident. Cells were frozen at -80oC overnight before being allowed to thaw at
room temperature (RT) for 1 h. The cells were subjected to a second freeze-thaw (3-5h) at -
80ºC and then centrifuged at 2500 rpm (Centurion) for 2.5 min to remove the cell debris. The
95
supernatant was filtered through a 0.2 µm Millex filter, divided into aliquots and stored at -
80oC. The viral titre was estimated by determination of TCID50 (Section 2.4).
2.4 50% Tissue Culture Infectious Dose (TCID50) Assay
TCID50 values for both FCV and MNV-1 were determined as follows: nine serial
dilutions (10-1 – 10-9) of stock virus were prepared in serum-free medium (MEM for FCV and
DMEM for MNV), 50 µl of each dilution were added to six wells of a 96-wells round-bottom
plate. One row was used as a control (no virus). Cells were prepared at a concentration of 4 x
105 cells/ml. One-hundred microliters of cell suspension were added to each well and the
cells were incubated at 37ºC with 5% CO2 overnight for FCV or 48 h for MNV. After
incubation, each well was scored as infected (positive) or uninfected (negative) and the titre
of the virus was calculated using the following formula from the method of Reed and
Muench.
Viral titre = [(% of wells infected at dilution above 50%) - 50%] .
[(% of wells infected above 50%) – (% of wells infected below 50%)]
2.5 Preparation of VPg-linked FCV RNA
Four 175-cm2 flasks containing of CRFK cells (80-100% confluent) in 30 ml medium
were used for infection with FCV. 20 ml of medium was removed from the cells and
collected in a sterile bottle. The cells were infected in the remaining 10 ml with FCV Urbana
(MOI=3) and left to rock for 30 min at RT. 5 ml of CRFK medium was added to the flask and
the cells were incubated for 6 hours at 37ºC with 5% CO2. The infected cells were scraped
off using cell scraper and decanted into 20 ml Universals. The cells were pelleted at low spin
speed (700 rpm) (Centurion) for 5 min. The supernatant was removed and filtered through a
0.2 µm Millex filter, aliquoted and stored at -80ºC. The cell pellets were pooled and washed
96
in 10 ml of PBS and spin at 2500 rpm (500g; Centurion) for 5 min. The cells were
resuspended in 4 ml of cold TN buffer (10 mM Tris [pH 7.8], 10 mM NaCl) and allowed to
swell on ice for 15 min. The cells were lysed with 60 strokes in a cold 2 ml tight-fitting glass
Dounce homogenizer (Bellco Glass, Inc. Vineland, N.J.). The lysate was transferred to four
1.5-ml Eppendorf tubes and centrifuged for 5 min at 3450 rpm (800g) at 4ºC (Micro 22R,
Hettich) to remove nuclei and un-lysed cells. The supernatants were then transferred to a
fresh Eppendorf tube and subjected to centrifugation for 20 min at 15000 rpm at 4ºC (Micro
22R, Hettich). The resulting pellets corresponding to replication complexes were each
resuspended in 120 µl of TN buffer containing 15% glycerol (replication complexes) and
stored at -80ºC. The replication complexes were used to make RNA at a later date.
2.6 Preparation of VPg-linked MNV RNA
RAW264.7 cells from (approximately 80–100 %) were infected with 1 ml of MNV-1
stock in 10 ml of FBS-Eagle’s DMEM and rocked for 1h. A further twenty ml of FBS-
Eagle’s DMEM were added to the flask and incubated for 23h at 37ºC with 5% CO2 until the
cytopathic effect was evident. After incubation the cells were scraped off and pelleted at 2500
rpm (Centurion) for 3 min. The supernatant was removed and filtered through a 0.2 µm
Millex filter, aliquotted and stored at -80ºC. The cell pellets were resuspended in 1 ml lysis
buffer supplemented with β-mercaptoethanol (GenElute kit Sigma).
2.7 Preparation and quantitation of VPg-linked FCV and MNV RNA
The GenElute Mammalian Total RNA Miniprep Kit (Sigma) was used to prepare
VPg-linked RNA from replication complexes as follows: 500µl of lysis buffer (2-ml lysis
buffer and 20µl 2-mercaptoethanol; prepared in the fume hood) was containing 250µl of
replication complexes mixed gently. The 750µl solution were then transferred into 4 blue
97
filter tubes and subjected to centrifugation at full speed for 1 min. The supernatant was
retained and equal volume of 70% ethanol was added prior to mixing. Lysate/ethanol was
applied to RNA-binding column in two aliquots and washed with 500µl of wash 1. The filter
of RNA-binding column was transferred into new tube and wash twice with 500µl of wash 2.
The columns were transferred into new tubes and 50µl of elution buffer was added. The tubes
were incubated at room temperature for a few minutes then spun at full speed (Boeco) for 1
min and the elution was collected and stored at -80ºC. One microliter of elution was used to
determine the concentration of the RNA using a Nano Drop ND-1000 spectrophotometer.
2.8 Immunofluorescence studies
2.8.1 Cell culture and virus infection
Cell of all types were maintained as previously described in Section 2.1. Cells were
seeded on 13-mm glass coverslips (VWR) at a required density overnight and virus infection
were carried out at a multiplicity of infection (MOI) of 0.2 for FCV or 30 for MNV at
different time points: 0, 2, 4, 6 and 8h and 0, 4, 8, 12, 16, 18 h, for FCV and MNV-1
respectively, unless otherwise specified. In all immunofluorescence studies, virus growth
medium was identical to the maintenance medium, except for the use of FBS at a
concentration of 2%. Mock-treated cells received 2% FBS growth medium only.
Optimization of MOI and concentration of PBS used were determined empirically during this
study.
2.8.2 Immunofluorescence assay
For immunofluorescence, cells were grown on 13-mm glass coverslips and following
chemical, physical treatment and infection, cells were fixed in 800 µl 4% paraformaldehyde
98
(Sigma Aldrich) in PBS (Appendix 1) on ice for 1h, then maintained in PBS at 4ºC until
staining. For staining, fixed cells were permeabilised by incubating with 0.1% Triton X-100
in PBS (BDH LTD) (Appendix 1) for 15 min, and then washed once with PBS. Fixed cells
were then blocked in PBS/BSA, PBS containing 1% bovine serum albumin (BSA) (Sigma
Aldrich) (Appendix 1) for 30 min. Primary and secondary antibodies (Table 2.1) were
diluted in blocking buffer. Following blocking, primary antibodies (200 µl) were added to
each well and incubated for 1 h, washed three times with PBS (3 x 5 minute washes), before
incubating for an additional 1 h with 200 µl secondary antibodies. A further 3X washes were
given before ToPro3 nuclear stain (1:10000 in PBS; Molecular Probes, Invitrogen) was
applied for 10 min. The coverslips were removed, rinsed in MQ water and placed cell-side
down on a glass slide with Vectashield mounting media (Vector Laboratories) then sealed
with clear nail varnish. All staining steps were performed at room temperature (RT). A Zeiss
LSM 510 META confocal laser microscope was used to detect SGs or PBs during infection
or chemical/physical treatment. The qualification of SGs and PBs formed were carried out
using the Image J software to determine the number of SG or PB-positive cells.
2.9 Effect of calicivirus infection on SGs and PBs formation
The MOI of the viruses used during this study was determined empirically using
confocal microscopy by calculating the percentage of infected cells as compared to the total
number of cells at different MOI. In addition, the dilutions of viral antibodies, SGs markers
proteins and PBs markers proteins used during this study were also selected empirically using
Confocal microscope (Table 2.1).
99
Figure 2.1 Primary and secondary antibodies that are used in
immunofluorescence studies
Primary antibody Company
and reference
number
Dilution Secondary antibodies Company
and reference
number
Dilution
Stress granules markers
G3BP1 BD
611126
1:600 Alexa-Anti-mouse 555 Invitrogen
A21424
1:400
PABP-1 Santa Cruz
Sc-32318
1:200 Alexa-Anti-mouse 555 Invitrogen
A21424
1:400
TIA-1 Santa Cruz
Sc-34739
and
Proteintech
17649-1-AP
12133-2-AP
Donkey anti-goat 555 Invitrogen
A21432
and
Santa Cruz
Sc-2024
1:400
eIF4G Santa Cruz
Sc-9601
Anti-mouse 555 Invitrogen
A21424
1:400
eIF3η Santa Cruz
Sc-16377
Donkey anti-goat 555 Invitrogen
A21432
1:400
P-bodies Markers
Dcp1 Santa Cruz
Sc-22574
1:50 Alexa-Anti-mouse 555 Invitrogen
A21424
1:400
Xrn-I Santa Cruz
Sc-165984
Alexa-Anti-mouse 555 Invitrogen
A21424
1:400
Viral markers
1G9 1:10000 Alexa-Anti-rabbit 488 Invitrogen
A11034
1:400
P76 1:600 Alexa-Anti-rabbit 488 Invitrogen
A11034
1:400
3D-polymerase
(NS7)
1:1500 Alexa-Anti-rabbit 488 Invitrogen
A11034
1:400
100
2.9.1 Staining for FCV-p76 protein
To visualise FCV infection, CRFK cells were stained with different dilution of anti
FCV p76 primary antibody (1:600; optimising during this study); Alexa-Fluor®-labelled anti-
rabbit secondary antibody (1:400; Molecular Probes, Invitrogen) and ToPro3 staining of
nuclei.
2.9.2 Staining for MNV-3D polymerase protein
To visualise MNV-1 infection, RAW264.7 cells were stained with different dilution
of anti MNV-3D-polymerase primary antibody (1:600; optimising during this study); Alexa-
Fluor®-labelled anti-rabbit secondary antibody (1:400; Molecular Probes, Invitrogen) and
ToPro3 staining of nuclei.
2.9.3 Staining for SG-markers
To visualise SGs formation during calicivirus infection, CRFK or RAW264.7 cells
were stained with different dilution of anti G3BP1, PABP-1, eIF4G, eIF3η, and TIA-1
primary antibodies; Alexa-Fluor®-labelled anti-mouse secondary antibody (1:400; Molecular
Probes, Invitrogen) for G3BP1, PABP, and eIF4G and donkey anti-goat IgG secondary
antibody (1:400; Molecular Probes, Invitrogen) for TIA-1 and eIF3ŋ. ToPro3 was used for
staining of nuclei.
101
2.9.4 Staining for PBs-markers
To visualise PBs formation during calicivirus infection, CRFK or RAW264.7 cells
were stained with different concentration of anti Dcp1 and Xrn1 primary antibodies; Alexa-
Fluor®-labelled anti-mouse secondary antibody (1:400; Molecular Probes, Invitrogen) and
ToPro3 staining of nuclei.
2.10 SGs formation during FCV infection
CRFK cells as well as FEA cells were plated on 13-mm coverslips slides (VWR) in
24 well plates at a concentration of 4 x 104 per well and maintained at 37°C with 5% CO2
overnight as described previously (Section 2.1.1 & 2.1.2). The next day, cells were infected
with 2% FBS MEM containing FCV at MOI of 0.2 at different time points (0, 2, 4, 6, and 8
h), the plates were rocked for 5 min and incubated at 37°C with 5% CO2 for 1 h. Mock-
treated cells received 2% FBS MEM only. Following incubation, viruses were removed and
replaced by 200 µl of 2% FBS MEM medium and the plates were incubated at 37°C with 5%
CO2 for additional 5 h. Cells were fixed for 1 h in 4% paraformaldehyde on ice for 1h, then
maintained in PBS at 4ºC until staining. The infected and mock infected cells were stained
for SGs markers (G3BP1, PABP-1, TIA-1, eIF3η and eIF4G) and p76 as a marker for virus
as described previously (Section 2.8.2).
2.10.1 Treatment with sodium arsenite (NaAsO2)
To select the optimal time of NaAsO2 (Sigma) (Appendix 1) treatment, CRFK cells
were grown on coverslips overnight prior treated the cells with 0.5mM NaAsO2 for different
times (30 min, 1h and 2h), then cell were fixed with 4% paraformaldehyde prior
102
immunostained for G3BP1 for detection the SGs using immunofluorescence microscopy as
described in Section 2.8.2.
2.10.2 Effect of FCV infection on SGs formation
CRFK and FEA cells were grown on 13-mm coverslips in 24-wells plate overnight
and then infected with FCV as above described for 6 h. Cells were then treated with 0.5 mM
NaAsO2 (Sigma) for 1h and then immunostained for G3BP1, PABP-1, TIA-1, eIF3η , eIF4G
and p76. Uninfected cells were treated with 0.5 mM NaAsO2 for 1h, infected cells as well as
untreated uninfected cells were used for comparison. The effect of FCV infection on SGs
formation was analysed using immunofluorescence microscopy as described in Section 2.8.2.
2.10.3 Effect of NaAsO2–induced SGs on FCV replication
CRFK cells were grown on 35-mm dishes overnight then treated with 0.5 mM
NaAsO2 for 1h, prior infected with FCV at different time points (0, 2, 4, 6, 8, 12, 16, and
18h). Cells supernatant then collected and used to determine the viral titre by TCID50
(Section 2.4) compared with supernatant of untreated infected cells. In addition, cell
supernatants at different time points were incubated with 13-mm coverslips containing CRFK
cells, fixed and immunostained for G3BP1 and p76 as described in Section 2.8.2.
2.10.4 Effect of FCV infection on hydrogen peroxide (H2O2) induced SGs
CRFK cells were grown on 13-mm coverslips and infected with FCV as above
described for 6 h. Cells were then treated with 1 mM H2O2 (Sigma) (Appendix 1) for 2h.
Uninfected cells were treated with 1 mM H2O2 for 2h, untreated cells and cell only infected
103
with FCV were used for comparison. Cells were stained for G3BP1 and p76 as a marker for
SGs and virus respectively. Immunofluorescence microscopy was used to determine the
effect of FCV infection on SGs formation as described in Section 2.8.2.
2.10.5 Effect of the H2O2-induced SGs on FCV replication
CRFK cells were plated on 35-mm dishes overnight, treated with 1 mM H2O2 for 2h,
and then infected with FCV at different time points (Section 2.8.1). Cells supernatant then
collected at each time point and used to determine the viral titre using TCID50 (Section 2.4).
In addition, cells at different time points were immunostained for G3BP1 and p76 to visualise
the SGs and virus as described in Section 2.8.2.
2.11 Preparation of FCV free supernatants
2.11.1 Virus Infection and clarification (virus elimination)
CRFK cells were seeded in 35-mm dishes at concentration of 2 x 105 cells and
incubated overnight. On the following day, the cells were infected with FCV for 6h, the
supernatant were then collected, transferred to Eppendorf tubes and spun at 4000 rpm at 4ºC
(Micro 22R, Hettich) for 30 min. After centrifugation, supernatant was filtered through a 0.2
µm Millex filter (Figure 2.2).
104
Figure 2.2 Preparation of the FCV-free supernatant from FCV-infected cells for
immunofluorescence studies.
105
2.11.2 Virus Precipitation
To precipitate the virus, solid NaCl was added to the supernatant at a final
concentration of 0.2M whilst mixing in cold room. Then, PEG3350 was slowly added to a
final concentration of 10%. The sample was then incubated onto a rotating wheel and mixed
overnight at 4ºC. The following morning, the samples were centrifuged for 60 minutes at
14000 rpm (Micro 22R, Hettich) at 4ºC. The supernatants were transferred to ultracentrifuge
tubes and span at 50000 rpm for 2h at 4ºC using SW55Ti rotor (Beckman). The supernatant
was then filtered through a 0.2 µm Millex filter, divided into aliquots (100µl each), and
stored in -20ºC until required.
2.13 Virus inactivation using ultraviolet-light (UV-light)
FCV pools or supernatant were UV inactivated at a wavelength of 254 nm for 4 min
(3 times) using a cross linker (Stratalinker® UV Crosslinker) before being used to inoculate a
wells of a 24-well plate containing CRFK cells. The cell were infected with FCV or FCV
supernatant and incubated at 37 ºC with 5% CO2 in a Galaxy S Incubator for 1 and 6 h. Cells
were then fixed using 4% paraformaldehyde for 1h prior to maintenance with PBS in 4ºC
until immunofluorescence study were carried out. In addition, the virus titre (with or without
UV treatment) were determined using TCID50 (Section 2.4) to confirm UV inactivation of the
virus.
106
2.14 Effect of cell culture supernatant on stress granules formation
CRFK, FEA, RAW264.7, BV2, 293T, J744, and THP-1 cells were grown on 13-mm
coverslips overnight before incubated for 6 h with the supernatant of FCV-infected or mock-
infected. Cells were then fixed and stained for G3BP1 and p76. Untreated CRFK cells were
incubated with solution of NaCl and PEG3350 at the same concentration as used in virus
precipitation protocol and followed by staining for G3BP1 to detect any effect on SGs
induction as described in Section 2.8.2.
2.15 Effect of interferon on stress granules formation
CRFK cells were grown on 13-mm coverslips overnight before incubating with
different concentration (150, 200, 300, and 3000 IU/ml) of interferon α (IFN α) for 6 h. Stress
granules were identified by immunofluorescence microscopy as described in Section 2.8.2.
2.16 Effect of RNAse A treatment on stress granules formation
RNAse A was added to FCV-infected CRFK supernatant (1U/1000 µl), incubated at
37oC for 15 min. CRFK cells were cultured on coverslips in 24-well plate overnight and then
treated with 200 µl of supernatant containing RNAse A. Control cells were incubated with
untreated supernatant. Cells were rocked for 10-15 min, incubated at 37ºC with 5% CO2 for 6
h, fixed using 4% paraformaldehyde for 1h and then immunostained for G3BP1 and p76 for
immunofluorescence microscopy as described in Section 2.8.2.
107
2.16 Effect of heat shock on stress granules formation
FCV-infected cells supernatant and mock-infected supernatant were incubated at 65ºC
for 10 and 20 min. 200µl of pre-heated supernatant was incubated with CRFK cells grown on
13-mm coverslips in 24-wells as previous described, rocked for 10-20 min and incubate at
37ºC with 5% CO2 for 6 h. Cells were then fixed with 4% paraformaldehyde for 1h and
stained for G3BP1 and p76 for immunofluorescence microscopy as described in Section
2.8.2.
2.17 SGs formation during MNV infection
RAW264.7 cells as well as BV-2 cells were cultured overnight as previously
mentioned (Section 2.1.2 &2.1.3). The following day, cells were plated on 13-mm coverslips
slides in 24-well plates at a concentration of 4.5 x 104 per well and incubated at 37°C with
5% CO2 overnight. The following day, cells were infected with MNV-1 at MOI of 30 for
different time (0, 4, 8, 12, 16 and 18 h). The plates were rocked for minutes and incubated at
37°C with 5% CO2 for 1 h. Mock-treated cells received media only. Following incubation,
the media were removed and replaced by 200 µl of 2% FBS MEM medium and the plates
were incubated at 37°C with 5% CO2. Cells were then fixed using 4% paraformaldehyde on
ice for 1h, then maintained in PBS at 4°C until staining. MNV-1 and mock infected cells
were stained for SGs markers (G3BP1, TIA-1, and eIF3η) and 3D-polymerase as a marker
for virus for immunofluorescence microscopy as described in Section 2.8.2.
108
2.18 Formation of P-bodies during FCV Infection.
CRFK and FEA Cells were seeded in 13-mm coverslips (VWR) in 24-well plates at a
concentration of 4 x 104 per well and maintained at 37°C with 5% CO2 overnight as
previously described. Next morning, cells were then infected with FCV at MOI 0.2 at
different time (0, 2, 4, 6 and 8h), cells then fixed after each time points with 4%
paraformaldehyde for 1h and maintained in PBS until staining for PBs markers. Cells were
then stained for Dcp1 and Xrn1 as PB markers as well as for p76 as virus marker for
immunofluorescence microscopy as described in Section 2.8.2.
2.19 Formation of P-bodies during MNV Infection.
RAW264.7 and BV2 cells were seeded in 13-mm coverslips (VWR) in 24-well plates
at a concentration of 4.5 x 104 per well and maintained at 37°C with 5% CO2 overnight as
previously described. Next morning, cells were infected with MNV-1 at MOI 30 for different
time (0, 4, 8, 12, and 18h), cells were then fixed after each time points with 4%
paraformaldehyde for 1h and maintained in PBS until staining for PB markers. Cells were
then stained for Dcp1and Xrn1 as PB markers as well as for 3D-polymerase as virus markers
for immunofluorescence microscopy as described in Section 2.8.2.
2.20 Statistical analysis
Experiments were carried out in triplicate with two samples each time, except where
specified. GraphPad Prism 6 was used for statistical testing using one-way ANOVA and
Dunnets post-doc testing to compare more than 2 samples or t-tests for comparing two
samples. Statistical significance was considered as P < 0.05. Image J software was used for
109
quantification of cells displayed stress granules or P-bodies during calicivirus infection. More
than 10 fields were randomly chosen and counted, with at least 400 cells counted for each
sample slide.
110
Chapter 3
Regulation of Stress Granule
formation during FCV and MNV-1
infection
111
3.1 Optimisation of experimental conditions for infection and SGs detection
To control the host response to infection and allow viral persistence, many viruses have
the ability to modulate the formation of cytoplasmic RNA granules in infected cells (Reineke
& Lloyd, 2013; Tsai & Lloyd, 2014). To date, the role played by RNA granules in the virus-
host relationship mediated by caliciviruses remains unknown. Therefore, to gain insights into
how caliciviruses modulate host stress responses during infection, I used FCV and MNV-1 as
surrogate models for human norovirus, and examined their effect on SG formation. As this was
the first project to investigate this line of research in our laboratory, a large body of my initial
work focussed on the optimisation of experimental conditions for immunofluorescence
detection of viruses and choice of marker proteins for the detection of SG assembly.
First, to determine the optimal MOI required for getting appropriate infection, I infected
the CRFK cells for 6h with different FCV MOI, from 0.1 to 1, and RAW264.7 cells for 12h
with different MNV MOI from 0.2 to 30. Cells were then fixed and stained for FCV VP1, the
capsid protein, or MNV NS7, the pol protein (see introduction, section 1.12), prior to indirect
immunofluorescence assay using confocal microscopy to determine the proportion of infected
cells (Figure 3.1 & 3.2). Among the different MOI tested, the best results were obtained at an
MOI of 0.2 for FCV and 30 for MNV where between 65-75% of the cells were found to be
infected (Figure 3.1C & 3.2K). These MOI were adopted for all subsequent experiments as
they allowed investigation of the formation of cytoplasmic granules in both infected and
neighbouring bystander cells during calicivirus infection.
Several dilutions of antibodies directed against p76, the pol-pro fusion protein, from
1:300 to 1:600 were then used to identify the optimal antibody concentration yielding strong
and specific labelling with low background noise in CRFK cells. As seen in Figure 3.3, the p76
112
dilution 1:600 gave an optimal signal: noise ratio for the detection of viral particles and this
dilution was then used in all subsequent studies.
113
ToPro-3
ToPro-3
ToPro-3
P76
P76
IG9
P76
Overlay
Overlay
Overlay
Figure 3.1 Optimization of FCV MOI for immunofluorescence studies. CRFK cells
were infected for 6h at several MOI as indicated (B-F), then cells were fixed and stained
for FCV infection using 1G9 antibody (green; middle panels) followed by Alexa-488
secondary antibody. ToPro3 stained nuclei are shown in blue (left panels). Right panels
show an overlay of both antibodies. Cells were visualised using a ZEISS LSM META
confocal laser microscope. The bar indicates a size of 10 µm.
A
B
C
D
E
F
- FCV
FCV
MO1 0.1
FCV
MOI 0.2
FCV
MOI 0.3
FCV
MOI 0.6
FCV
MOI 1
1G9
ToPro-3 1G9 Overlay
ToPro-3 1G9 Overlay
1G9
ToPro-3 Overlay 1G9
114
-MNV-1
MNV-1
MOI 0.2
MNV-1
MOI 0.5
MNV-1
MOI 1
MNV-1
MOI 2
MNV-1
MOI 5
MNV-1
MOI 10
A
B
C
D
E
F
G
ToPro-3 Overlay NS7
ToPro-3 Overlay NS7
Overlay NS7 ToPro-3
Overlay NS7 ToPro-3
Overlay NS7 ToPro-3
Overlay NS7 ToPro-3
Overlay NS7 ToPro-3
- Continued -
115
Figure 3.2 Optimization of MNV-1 MOI for immunofluorescence studies.
RAW cells were infected for 12h at several MOI as indicated (B-K), then cells
were fixed and stained for MNV-1 infection using NS7 antibody (green, middle
panels) followed by Alexa-488 secondary antibody. ToPro3 stained nuclei are
shown in blue (left panels) and the right panels show an overlay of both antibodies.
Cells were visualised using a ZEISS LSM META confocal laser microscope. The
bar indicates a size of 10 µm.
MNV-1
MOI 15
MNV-1
MOI 20
MNV-1
MOI 25
MNV-1
MOI 30
H
I
J
K
ToPro-3 Overlay NS7
ToPro-3 Overlay NS7
ToPro-3 Overlay NS7
ToPro-3 Overlay NS7
116
Figure 3.3 Optimization of p76 antibody dilution for immunofluorescence
studies. CRFK cells were infected for 6h at MOI of 0.2, then infected (B-E) and
uninfected (A) cells were fixed and stained for FCV infection using several dilution
of p76 antibody (B-D) followed by Alexa-488 secondary antibody (green). ToPro-3
stained nuclei are shown in blue (left panels) and middle panels show p76 while the
right panels show an overlay of both antibodies. Cells were visualised using a ZEISS
LSM META confocal laser microscope. The bar indicates a size of 10 µm.
ToPro-3
ToPro-3
ToPro-3
ToPro-3
p76
p76
p76
p76
Overlay
Overlay
Overlay
Overlay
A
B
C
D
E
- FCV
FCV
1:300
FCV
1:400
FCV
1:500
FCV
1:600
Overlay p76 ToPro-3
117
3.2 Chemical induction of SGs using oxidative stress in CRFK cells
SG formation is induced by a wide range of experimental stresses such as heat shock
(Anderson & Kedersha, 2002; Kedersha et al., 1999; Nover et al., 1983), UV irradiation
(Anderson & Kedersha, 2006; Moutaoufik et al., 2014), hypoxia(Anderson & Kedersha, 2006;
Takahashi et al., 2013), exposure to sodium arsenite (Anderson & Kedersha, 2006; Khaperskyy
et al., 2014), hydrogen peroxide (Emara et al., 2012) and Pateamine A (Anderson & Kedersha,
2009b; Dang et al., 2006; Khaperskyy et al., 2014). These are useful tools to dissect whether
stress granules are assembled in response to stresses communicated in an eIF2-dependent or
independent manner.
The most commonly used SG inducer is sodium arsenite treatment which results in the
phosphorylation of eIF2α via the HRI kinase (Anderson & Kedersha, 2009b; Kedersha &
Anderson, 2007; Kedersha et al., 1999; McEwen et al., 2005). To assay the ability of CRFK
cells, supporting FCV replication, to respond to stress and form SGs, we treated these cells
with 0.5 mM of sodium arsenite for 30 min, 1h or 2h and subsequently monitored the assembly
of SGs using G3BP1 as marker (Figure 3.4). The immunofluorescence analysis revealed that
1h was the optimal length of treatment for induction of SGs, reflected in the strong
accumulation of cytoplasmic G3BP1 foci, while 30 min did not result in any foci formation,
and 2h led to abnormal morphology and death of most of the cells (Figure 3.4D).
118
Figure 3.4 Induction of stress granules by oxidative stress. CRFK cells were
incubated with 0.5 mM sodium arsenite (SA) for various time points as mediated
on the panels B-D, then cells were fixed and stained for G3BP1 antibody (red;
middle panels), followed by Alexa-555 secondary antibody. ToPro-3 stained
nuclei are shown in blue. Cells were visualised using a ZEISS LSM META
confocal laser microscope. The bar indicates a size of 10 µm.
A
B
C
D
- SA
+ SA
30 min
+ SA
1h
+ SA
2h
ToPro-3 G3BP1 Overlay
ToPro-3 G3BP1 Overlay
ToPro-3 G3BP1 Overlay
ToPro-3 G3BP-1 Overlay
119
3.3 Optimisation of the markers and conditions used for stress granule detection in CRFK
cells
To monitor SG assembly during infection and not rely on only one marker, G3BP1, we
assayed the potential use of diverse SG markers in CRFK cells. Based on informal discussion
with Dr Dominique Weill in Université Paris 6 (Paris, France), we opted for the following
potential stress granule markers: G3BP1, TIA-1, PABP, eIF4G, eIF3. CRFK cells were
stressed with sodium arsenite (0.5 mM 1h) and the assembly of SGs was monitored by
immunofluorescence microscopy using G3BP1, PABP, TIA-1, eIF3η and eIF4G, using
unstressed cells as control. While sodium arsenite treatment resulted in the accumulation of
G3BP1 and PABP-1 cytoplasmic foci (Figure 3.5 A & B), no signal could be detected for
eIF4G, eIF3η and TIA-1 (Figure 3.5C, D & E). For all antibodies, a large range of dilutions
was used to identify the most suitable dilution for SG detection (data not shown).
120
eIF4G eIF4G
eIF3η
TIA-1
TIA-1
eIF3η
G3BP1 G3BP1
PABP-1 PABP-1
eIF3η
TIA-1
G3BP1
Figure 3.5 Detection of stress granules by oxidative stress in FCV-infected cells
using various markers. CRFK cells were incubated with 0.5 mM sodium arsenite (SA)
for 1h, treated and untreated cells were then fixed and stained for (A) G3BP1, (B)
PABP-1, (C) eIF4G or (D) eIF3η and (E) TIA-1 followed by Alexa-555 secondary
antibody. ToPro3 stained nuclei are shown in blue. Cells were visualised using a ZEISS
LSM META confocal laser microscope. The bar indicates a size of 10 µm.
- SA + SA
A
B
C
D
E
G3BP1
121
The amount of serum used in the cell culture media is a key component of a successful
infection but can also act as a source of stress on cells. For instance, serum-starvation is a potent
inducer of SG formation in certain cell types used for cell culture systems (Anderson &
Kedersha, 2009b; Dewey et al., 2011; Kedersha et al., 1999). The use of serum and glucose-
free medium can induce SG assembly in HepG2 hepatocytes cells (Hanson & Mair, 2014).
The common protocol for FCV infection recommends the use of serum-free media,
which could potentially induce stress, therefore I analyzed the effect of fetal bovine serum
(FBS) concentration on SG assembly by incubating CRFK cells with different concentrations
of FBS (0, 2, 5 and 10%) and monitored the impact on SGs by immunofluorescence microscopy
using G3BP1 as SG marker (Figure 3.6). While the absence of serum induced the formation of
stress granules, the lowest FBS concentration tested, 2%, did not and was selected for the
subsequent infections.
122
ToPro-3 G3BP1 Overlay
ToPro-3 G3BP1 Overlay
ToPro-3 G3BP1 Overlay
ToPro-3 G3BP1
0% FBS
2% FBS
5% FBS
10% FBS
A
B
C
D
Overlay
Figure 3.6 Optimization of FBS concentration for detect of stress granules in CRFK
cells. CRFK cells were grown overnight using different concentration of FBS as indicated
(A-D), cells were then fixed and stained for G3BP1 antibody (red; middle panels), followed
by Alexa-555 secondary antibody. ToPro3 stained nuclei are shown in blue. Cells were
visualised using a ZEISS LSM META confocal laser microscope. The bar indicates a size
of 10 µm.
123
3.4 Optimisation of the markers used for stress granule detection in RAW264.7 cells
To monitor SG assembly in a cell line supporting MNV-1 infection, we stressed
RAW264.7 cells with sodium arsenite (0.5 mM 1h) and used immunofluorescence microscopy
to detect G3BP1, TIA-1 and eIF3, using unstressed cells as control. Unexpectedly, none of
the above markers displayed aggregation into cytoplasmic foci following sodium arsenite
treatment (Figure 3.7). Several different antibody dilutions were tested without any further
success (not shown). These results suggest that RAW264.7 may be intrinsically unable to
assemble SGs, at least in response to oxidative stress.
124
G3BP1
TIA-1
eIF3η eIF3η
TIA-1
G3BP1
Figure 3.7 Detection of stress granules in RAW264.7 cells using various markers.
RAW264.7 cells were treated with 0.5mM SA , then fixed and stained for (A) G3BP1 or
(B) TIA-1 and (C) eIF3η followed by Alexa-555 secondary antibody. ToPro3 stained nuclei
are shown in blue. Cells were visualised using a ZEISS LSM META confocal laser
microscope. The bar indicates a size of 10 µm.
- SA + SA
A
B
C
125
3.5 Stress granule formation during FCV infection.
To investigate the effect of FCV infection on the formation of SGs and whether
cytoplasmic granules are part of the response to the virus, CRFK cells were infected with FCV
Urbana at MOI 0.2 for 8h, and the presence of SGs was investigated by immunofluorescence
assay microscopy at several time points: 0h, 2h, 4h, 6h and 8h using G3BP1 as marker for
stress granules and p76 as marker for viral particles (Figure 3.8). For each time point in the
FCV-infected or mock-infected cells, the number of cells displaying SGs were quantified and
the experiments performed in triplicate (Figure 3.9).
The experiments revealed that in mock-infected cells only very few cells were able to
form SGs over the 8h time course. Overall, SGs were only detected at 4, 6 and 8h post-infection
(Figure 3.14 panels C, D and E). The induction was modest was a maximum of 15% of cells
displaying SGs at 4h post-infection, against 1.4% for mock-infected cells. A closer inspection
of the results actually indicates that although overall the infection results in an increase in the
number of cells displaying SGs, we could not detect any SGs in cells infected with FCV (see
Figure 3.8D for a close up view at 6h post-infection). All SGs formed are detected in
uninfected, neighbouring or bystander cells that are in the vicinity of infected cells (Figure 3.8).
In summary these experiments suggest that FCV infection does not induce SG formation in
infected cells, while SG formation was induced in uninfected cells located near foci of
infection.
126
Figure 3.8 Detection of stress granules during FCV infection. CRFK cells were infected
with FCV at MOI of 0.2 for several time points as mediated on the panels B-E, and then
immunostained for G3BP1 antibody (red) and p76 (green) as markers for stress granules
and virus respectively followed by Alexa-555 and Alexa-488 secondary antibodies. Left
panels show p76 alone (green), middle panels show G3BP1 alone (red) and right panels
show an overlay of both. Cell nuclei were stained with ToPro-3 (blue). On panel D a close-
up view is displayed on the side. Cells were visualised using a ZEISS LSM META confocal
laser microscope. The bar indicates a size of 10 µm.
p76 G3BP1 Overlay
p76 G3BP1 Overlay
p76 G3BP1 Overlay
p76 G3BP1 Overlay
A
B
C
D
E
- FCV
FCV 2hpi
2HPI
FCV 4hpi
2HPI
FCV 6hpi
2HPI
FCV 8hpi
2HPI
Overlay G3BP1 p76
127
h o u rs p o s t- in fe c t io n
Ce
lls
dis
pla
yin
g
str
es
s g
ra
nu
les
(%
)
0 2 4 6 8
0
5
1 0
1 5
2 0
2 5
- F C V
+ F C V
ns
ns
**
****
Figure 3.9 Quantification of the number of cells displaying stress granules during FCV
infection. CRFK were infected with FCV at MOI of 0.2 for several time points, stained for
G3BP1 and p76 as described in Figure 3.8. Confocal images were then used to quantify the
numbers of cells with stress granules using Image J software. Results shown are the
percentage of cells displayed SGs at each time point compared to the control, and are the
mean of three independent experiments (+/-SE; ** is P ≤0.01; ns not significant).
128
To confirm these results, the 6h time point was repeated using PABP-1 as a marker for
SGs. As shown in Figure 3.10, the infection also resulted in the induction of stress granules,
with 8% of cells displaying SGs during infection as compared to 3.9% for mock infected, but
this difference was not found to be significant after statistical analysis (Figure 3.11). However,
SGs were only detected in neighbouring cells close to infection foci and none could be detected
within FCV-infected cells (Figure 3.10).
Furthermore, to confirm that this effect was not cell-line specific, we infected feline
embryonic fibroblast (FEA) cells, that support FCV replication, with FCV for 6h and monitored
SG assembly using G3BP1 as marker for SGs and p76 as marker for the virus. Similarly to
results obtained with CRFK cells, infection of FEA cells with FCV did not result in SG
induction, and only uninfected cells displayed SGs (Figure 3.12).
129
PABP-1
PABP-1 p76
p76 Overlay
Overlay
Figure 3.10 Detection of stress granules during FCV infection using PABP-1 as a
marker. CRFK cells were infected with FCV at a MOI of 0.2 for 6h, then infected and mock
cells immunostained for PABP-1 (red) and p76 (green) as markers for stress granules and
virus respectively followed by Alexa-555 and Alexa-488 secondary antibodies. From the
left, the second panels show p76 alone (green), third panels show PABP-1 alone (red) and
right panels show an overlay of both. Left panels show cell nuclei were stained with ToPro-
3 (blue). Cells were visualised using a ZEISS LSM META confocal laser microscope. The
bar indicates a size of 10 µm.
- FCV
+ FCV
6hpi
130
- F
C V
+ F
CV
6h
0
5
1 0
1 5
2 0
2 5C
ell
s d
isp
lay
ing
str
es
s g
ra
nu
les
(%
)
n s
Figure 3.11 Quantification of the number of cells displaying stress granules during
FCV infection using PABP-1. CRFK were infected with FCV at MOI of 0.2 for 6h, stained
for PABP-1 and p76 as described in Figure 3.10. Confocal images were then used to quantify
the numbers of cells with stress granules using Image J software. Results show are the
percentage of cells displayed SGs at compared to the control, and are the mean of three
independent experiments (+/-SE; ns: not significant).
131
p76 G3BP1 Overlay
p76 G3BP1 Overlay
Figure 3.12 Detection of SGs in FEA cells during FCV infection. FEA cells were infected
with FCV at MOI of 0.2 for 6h, and then immunostained for G3BP1 antibody (red) and p76
(green) as markers for SG and virus respectively. Left panels show p76 alone (green), middle
panels show G3BP1 alone (red) and right panels show an overlay of both. Cell nuclei were
stained with ToPro-3 (blue). Cells were visualised using a ZEISS LSM META confocal laser
microscope. The bar indicates a size of 10 µm.
- FCV
+ FCV
6hpi
132
The previous optimisation section revealed that none of the following markers for SGs,
TIA-1, eIF4G or eIF3 allowed successful detection of SGs in unstressed cells. However, we
cannot exclude that this marker could accumulate in the granules detected during infection, in
neighbouring cells. To test for this, CRFK cells infected with FCV for 4h, which displayed SGs
as revealed using G3BP1, were assayed by immunofluorescence microscopy using TIA-1 or
eIF3 as stress granule markers. As shown in Figure 3.13, no cytoplasmic foci could be
detected with these markers, confirming that they do not allow successful detection of SGs in
CRFK cells, infected or stressed with sodium arsenite.
133
ToPro-3 p76 Overlay TIA-1
ToPro
-3 p76 Overlay TIA-1
p76 eIF3η overlay ToPro-3
p76 eIF3η overlay ToPro-3
p76 eIF4G overlay ToPro-3
A
B
+ FCV
p76 eIF4G overlay ToPro-3
- FCV
- FCV
+ FCV
- FCV
+ FCV
Figure 3.13 Detection of SGs formation during FCV infection using different markers.
CRFK cells were infected for 6h at MOI of 0.2, then uninfected and infected cells were
fixed and stained for FCV infection using p76 antibody (green) and for different SG marker
including (A) TIA-1, (B) eIF3η and (C) eIF4G antibody (red) followed by Alexa-488 and
Alexa-555 secondary antibodies. ToPro3 stained nuclei are shown in blue. Cells were
visualised using a ZEISS LSM META confocal laser microscope. The bar indicates a size
of 10 µm.
C
134
3.6 Stress granule formation during MNV-1 infection.
To investigate whether the absence of SGs was a pattern that was conserved in other
caliciviruses, I examined the formation of SGs during MNV-1 infection. While RAW264.7
cells do not form SGs in response to sodium arsenite treatment, we could not exclude that they
would respond to stress mediated by MNV-1 infection. To test this, RAW264.7 cells were
infected with MNV-1 at an MOI of 30 for 18h, and examined by immunofluorescence
microscopy at 0, 4, 8, 12 and 18h post-infection using G3BP-1 as a marker for SGs, and NS7
as marker for MNV-1 (Figure 3.14). MNV-1 infection did not result in the assembly of
cytoplasmic G3BP-1 foci at any of these times (Figure 3.14).
To confirm that this complete absence of SGs, unlike during FCV infection, was not
due to a cell-specific or marker-specific effect, RAW264.7 cells were also examined using
TIA-1 and eIF3η as SG markers at 12h post-infection (Figure 3.15). MNV-1 infection did not
result in assembly of cytoplasmic foci for any of these markers (Figure 3.15). Next, BV2
murine microglial cells, also supporting MNV-1 infection, were infected with MNV-1 for 12h
or stressed with sodium arsenite (0.5 mM 1h) and examined using G3BP1, TIA-1 and eIF3η
as SG markers (Figure 3.16). None of these experimental conditions led to the detection of SGs
during infection, and only G3BP1 accumulated in SGs following sodium arsenite stress.
Overall, this lead to the conclusion that cells that support MNV-1 infection do not
respond to sodium arsenite treatment or infection by assembling canonical stress granules and
therefore subsequent studies focussed only on the effect of FCV.
135
- SA/- MNV-1
+ SA/- MNV-1
+ MNV-1
0hpi
+ MNV-1
4hpi
+ MNV-1
8hpi
+ MNV-1
12hpi
+ MNV-1
18hpi
G3BP1 NS7 Overlay
G3BP1 NS7 Overlay
G3BP1 NS7 Overlay
G3BP1 NS7 Overlay
G3BP1 NS7 Overlay
G3BP1 NS7 Overlay
G3BP1 Overlay
Figure 3.14 Detection of stress granules during MNV-1 infection. RAW264.7 cells were infected
with MNV-1 at a MOI of 30 for different time points or treated with sodium arsenite (SA) as indicated
(B-G) and then immunostained for G3BP1 (red) and NS7 (green) as markers for stress granules and
virus respectively, followed by Alexa-555 and Alexa-488 secondary antibodies. Left panels show
NS7 alone (green), middle panels show G3BP alone (red) and right panels show an overlay of both.
Cell nuclei were stained with ToPro-3 (blue). Cells were visualised using a ZEISS LSM META
confocal laser microscope. The bar indicates a size of 10 µm.
NS7
A
B
C
D
E
F
G
136
- MNV-1
- MNV-1
+ MNV-1
+ MNV-1
TIA-1
eIF3η
Figure 3.15 Detection of stress granules during MNV-1 infection using various markers.
RAW264.7 cells were infected with MNV-1 for 12h, infected and mock cells were then fixed and
stained for (A) TIA-1 and (B) eIF3η followed by donkey anti goat secondary antibody. ToPro3 stained
nuclei are shown in blue. Cells were visualised using a ZEISS LSM META confocal laser microscope.
The bar indicates a size of 10 µm.
Overlay TIA-1 p76
Overlay TIA-1 p76
Overlay eIF3η p76
Overlay eIF3η p76
A
B
137
eIF3η
TIA-1
- SA/- MNV-1
+ SA/- MNV-1
+ MNV-1
12hpi
A
B
eIF3η Overlay p76
eIF3η Overlay p76
eIF3η Overlay p76
TIA-1 Overlay p76
TIA-1 Overlay p76
p76 TIA-1 Overlay
- SA/- MNV-1
+ SA/- MNV-1
+ MNV-1
12hpi
138
G3BP1
Figure 3.16 Detection of stress granules during MNV-1 infection in BV-2 cells using different
markers. BV-2 cells were infected with MNV-1 at a MOI of 30 for 12h or treated with sodium
arsenite (SA; 1h 0.5mM) and then immunostained for various markers for SG (A-C), and for MNV
infection using NS7 antibody. Left panels show NS7 alone (green), middle panels show G3BP1
alone (red) and right panels show an overlay of both. Cell nuclei were stained with ToPro-3 (blue).
Cells were visualised using a ZEISS LSM META confocal laser microscope. The bar indicates a
size of 10 µm.
C
Overlay G3BP1 p76
Overlay G3BP1 p76
Overlay G3BP1 p76
- SA/- MNV-1
+ SA/- MNV-1
+ MNV-1
12hpi
139
3.7 FCV infection impairs the assembly of stress granules induced by sodium arsenite.
It is well documented that the detection of viral dsRNA, formed as intermediate during
viral RNA replication, is a major stress for the host cell and a potent activator of stress pathways
resulting in SG formation (Dewey et al., 2011; Okonski & Samuel, 2013; Reineke & Lloyd,
2013; Takahashi et al., 2013; Zhang et al., 2014). The previous results in Figure 3.8, suggested
that FCV infection might actively prevent the formation of SGs during infection.
Therefore to dissect this further, we analysed whether FCV-infection could prevent the
formation of SGs induced by sodium arsenite. To this end, CRFK cells were either mock or
FCV-infected for 2h or 6h, and then were treated with 0.5 mM sodium arsenite for 1h to induce
the formation of SGs. The ability of CRFK cells to form SGs was then addressed by
immunofluorescence using G3BP1 as the SG marker. As shown on Figure 3.17, mock-infected
cells that are treated with sodium arsenite are competent in assembling SGs with 83.4% of cells
displaying G3BP1 foci (Figure 3.17, Figure 3.18). However, when cells were FCV-infected
prior to treatment with sodium arsenite the ability to assemble SGs was impaired with only 7%
of cells containing G3BP1 foci at 6h post-infection (Figure 3.17, Figure 3.18). In contrast, at
2h post-infection there is less effect of FCV infection on the ability of CRFK cells to assemble
SGs with 41% of cells displaying SGs (Figure 3.18). These results strongly suggest that FCV-
infection impairs the ability of cells to assemble stress granules in response to sodium arsenite,
and that this effect is more pronounced later in the replication cycle.
140
overlay G3BP1 p76
overlay G3BP1 p76
Figure 3.17 Detection of stress granules induced by NaAsO2 following FCV infection. CRFK cells were infected with FCV for 2h and 6h, treated with sodium arsenite (SA) to induce stress granule formation after each time for 1h and then immunostained for G3BP1 (red) and p76 (green) as markers for stress granule and virus respectively, followed by Alexa-555 and Alexa-488 secondary. Left panels show p76 alone (green), middle panels show G3BP alone (red) and right panels show an overlay of both. Cell nuclei were stained with ToPro-3 (blue). Cells were visualised using a ZEISS LSM META confocal laser microscope. The bar indicates a size of 10 µm.
-FCV/-SA
-FCV/+SA
+FCV 6hpi/-SA
+FCV 6hpi/+SA
overlay p76 G3BP1
overlay G3BP1 p76
overlay
overlay G3BP1 p76
G3BP1 p76
+FCV 2hpi/-SA
+FCV 2hpi/+SA
overlay
overlay
G3BP1 p76
p76
141
Ce
lls
dis
pla
yin
g
str
es
s g
ran
ule
s (
%)
-FC
V/-
SA
-FC
V/+
SA
+F
CV
2h
/-S
A
+F
CV
2h
/+S
A
+F
CV
6h
/-S
A
+F
CV
6h
/+S
A
0
2 0
4 0
6 0
8 0
1 0 0*
**
Figure 3.18 Quantification of the number of cells displaying stress granules induced
by NaAsO2 following FCV infection at different time points. CRFK were infected with
FCV at MOI of 0.2 for 2h and 6h, stained for G3BP1 and p76 as described in Figure 3.17.
Confocal images were then used to quantify the numbers of cells with stress granules using
Image J software. Results shown are the percentage of cells displaying SGs at each time
point compared to the control, and are the mean of three independent experiments (+/-SE;
* is P ≤0.05, ** is P≤0.01 for 2h and 6h respectively).
142
We further confirmed these results by using another SG marker, PABP, and the same
experimental design. As shown on Figure 3.19, mock-infected cells respond to sodium arsenite
treatment with 71.7% of cells assembling PABP cytoplasmic foci (Figure 3.20). In contrast,
only 21% of FCV-infected cells that are treated with sodium arsenite could assemble SGs
(Figure 3.19, Figure 3.20). Although the inhibition of SG induction was not as strong as when
using G3BP1 as marker, this confirms that FCV infection impairs the ability of cells to
assemble SGs in response to oxidative stress induced by sodium arsenite.
143
overlay PABP-1 p76
overlay PABP-1 p76
A
B
C
D
overlay PABP-1 p76
overlay PABP-1 p76
-SA/-FCV
+SA/-FCV
-SA/+FCV 6hpi
+SA/+FCV 6hpi
Figure 3.19 Detection of stress granules induced by sodium arsenite (SA) following
FCV infection using PABP-1 as a marker. CRFK cells were infected with FCV for 6h,
treated with sodium arsenite (SA; 0.5mM 1h) to induce stress granule formation and then
immunostained for PABP-1 (red) and p76 (green) as markers for stress granule and virus
respectively, followed by Alexa-555 and Alexa-488 secondary. Left panels show p76 alone
(green), middle panels show PABP-1 alone (red) and right panels show an overlay of both.
Cell nuclei were stained with ToPro-3 (blue). Cells were visualised using a ZEISS LSM
META confocal laser microscope. The bar indicates a size of 10 µm.
144
- F
CV
-S
A
- F
CV
+S
A
+ F
CV
-S
A
+ F
CV
+S
A
0
2 0
4 0
6 0
8 0
1 0 0
Ce
lls
dis
pla
yin
g
PA
BP
-1 c
on
tain
ing
-
SG
s (
%)
****
Figure 3.20 Quantification of the number of cells displaying stress granules induced by
sodium arsenite following FCV infection using PABP-1 as a marker. CRFK were
infected with FCV at MOI of 0.2 for 6hpi, stained PABP-1 and p76 as described in Figure
3.19. Confocal images were then used to quantify the numbers of cells with stress granules
using image J software. Results shown are the percentage of cells displayed SGs compared
to the control, and are the mean of three independent experiments (+/-SE; **** is P
≤0.0001).
145
3.8 FCV infection impairs the assembly of stress granules induced by hydrogen peroxide.
Several stress pathways are responsible for the induction of SGs in cells (Anderson &
Kedersha, 2009a; b; Mazroui et al., 2006; Montero & Trujillo-Alonso, 2011; Tsai & Lloyd,
2014). The results from section 3.7 suggest that FCV infection prevents the induction of SGs
that are mediated in an eIF2 phosphorylation dependent manner. SG assembly can however
also be induced by an eIF2 phosphorylation independent pathway (Dang et al., 2006; Emara
et al., 2012; Mazroui et al., 2006). For example, the induction of SGs by Pateamine A and
hippuristanol is not dependent on the phosphorylation of eIF2 but instead occurs through an
effect on the function of the RNA helicase eIF4A, impairing eIF4F activity and translation
(Dang et al., 2006; Mazroui et al., 2006). Furthermore, the induction of SGs by hydrogen
peroxide is mediated by the inhibition of the cap-binding eIF4F complex rather than through
the phosphorylation of eIF2α. Hydrogen peroxide treatment promotes the hypo-
phosphorylation of 4E-BP1 resulting in the sequestration of eIF4E and disruption of eIF4F
assembly (Emara et al., 2012).
Therefore to assess whether FCV infection could prevent the induction of SGs in an
eIF2 phosphorylation independent manner, we used hydrogen peroxide as a SG inducer.
CRFK cells were either mock or FCV-infected for 6h, and then were treated with 1 mM
hydrogen peroxide for 2h to induce the formation of SGs. The ability of CRFK cells to form
SGs was then analysed by immunofluorescence using G3BP1 as the SG marker. As shown in
Figure 3.21, mock-infected cells that are treated with hydrogen peroxide are competent in
assembling SGs with 87.7% of cells displaying G3BP1 foci (Figure 3.21, Figure 3.22).
However, when cells were FCV-infected prior to treatment with hydrogen peroxide, the ability
to assemble SGs was impaired with only 7.7% of cells containing G3BP1 foci (Figure 3.21,
146
Figure 3.22). These results strongly suggest that FCV infection impairs the ability of cells to
assemble stress granules in response to hydrogen peroxide and therefore in an eIF2
phosphorylation independent manner.
147
-FCV/-H2O2
-FCV/+H2O2
+FCV/-H2O2
+FCV/+H2O2
overlay G3BP1 p76
overlay G3BP1 p76
overlay G3BP1 p76
overlay G3BP1 p76
Figure 3.21 Detection of stress granules induced by hydrogen peroxide (H2O2) following FCV infection. CRFK cells were infected with FCV for 6h, treated with hydrogen (H2O2) to induce stress granule formation and then immunostained for G3BP1 antibody (red) and p76 antibody (green) as markers for stress granule and virus respectively, followed by Alexa-555 and Alexa-488 secondary. Left panels show p76 alone (green), middle panels show G3BP1 alone (red) and right panels show an overlay of both. Cell nuclei were stained with ToPro-3 (blue). Cells were visualised using a ZEISS LSM META confocal laser microscope. The bar indicates a size of 10 µm.
148
- F
CV
- H
2O
2
+ H
20
2
+ F
CV
+ F
CV
+ H
20
2
0
2 0
4 0
6 0
8 0
1 0 0
Ce
lls
dis
pla
yin
g
str
es
s g
ran
ule
s (
%)
***
Figure 3.22 Quantification of the number of cells displaying stress granules induced by
hydrogen peroxide (H2O2). CRFK were infected with FCV at MOI of 0.2 for 6hpi, stained
for G3BP1 and p76 as described in Figure 3.21. Confocal images were then used to quantify
the numbers of cells with stress granules using image J software. Results shown are the
percentage of cells displayed SGs compared to the control, and are the mean of three
independent experiments (+/-SE; *** is P ≤0.001).
149
3.9 Effect of Stress granule induction on FCV replication
Overall, FCV infection prevents the assembly of SGs induced by stress signalling via
both the eIF2-dependent and independent pathways. This raises the possibility that FCV
replication impairs the assembly of SGs to prevent the formation of intracellular compartments
that would putatively have an antiviral effect on the propagation of the virus.
To test this hypothesis, we monitored whether chemical induction of stress granules,
prior to infection with FCV, would alter viral replication. To this end, CRFK cells were treated
with either sodium arsenite or hydrogen peroxide to induce SG formation and the replication
of FCV was determined by measuring viral titres at 6 or 8h post-infection using a TCID50 assay.
As shown in Figure 3.23, the induction of SGs, either with sodium arsenite or hydrogen
peroxide impaired viral replication with reduction in titre from 1,61.10+6 to 0 TCID50/ml for
both treatment at 6h post-infection, and 6,21.10+6 to 0 TCID50/ml at 8h post-infection, for both
treatments. Therefore FCV infection may impair SG formation to prevent an antiviral response
from being triggered.
150
6h u
ntreate
d
6h S
A
6h H
2O
2
8h u
ntreate
d
8h S
A
8h H
2O
2
1 0 -1
1 0 0
1 0 1
1 0 2
1 0 3
1 0 4
1 0 5
1 0 6
1 0 7
TC
ID5
0/m
L
F C V in fe c t io n (h p i)
Figure 3.23 Effect of stress granules induction on FCV replication. CRFK cells were
treated with 0.5 mM sodium arsenite (SA) for 1h or 1 mM hydrogen peroxide (H2O2) for
2h, then infected with FCV at MOI of 0.2 for 6h (black) or 8h (grey). The supernatants of
infected cells and treated infected cells for each time-point were used to determination of
viral titre by TCID50. Results show viral titre from treated cells in each time point compared
to the viral titre from untreated infected cells, and are the mean of three independent
experiments.
151
3.10 Effect of FCV infection on pre-assembled stress granules
The results from sections 3.4, 3.7 and 3.8 strongly suggest that FCV infection is actively
impairing the formation of SGs as a response to various stresses. However, we also asked the
question whether FCV infection could induce the dissolution of pre-existing SGs. Upon
treatment with sodium arsenite, cells have the ability to assemble SGs. Following removal of
the stress, cells then recover while SGs are dissolved and many factors are suggested to be
involved in this process such as turnover of SG protein component, chaperone proteins, and
autophagy (Buchan, 2014). In addition, tyrosine-phosphorylation-regulated kinase 3 (DYRK3)
plays a possible role in stress granule dissolution through control of mTORC1 signalling during
cellular stress (Wippich et al., 2013). Furthermore, the disassembly of SGs upon recovery may
be due to dephosphorylation of eIF2α (Kedersha et al., 1999).
Therefore to test the ability of FCV infection to dissolve pre-existing SGs, CRFK cells
were treated with sodium arsenite for one hour then mock- or FCV-infected and the number of
cells displaying stress granules monitored over an 8h time course (Figure 3.24). Following the
removal of sodium arsenite, mock-infected cells quickly recovered from the stress, with the
number of cells displaying SGs dropping from 83% to 65% by 2h post-infection, then 32% by
4h post-infection and continuously dropping further. At similar times during FCV infection,
the number of cells displaying SGs were 20.8% and 43.2% at 0 and 2h post-infection
respectively and no stress granules could be detected at later time points (Figure 3.25).
Therefore this could suggest that pre-assembled SGs disappeared more quickly during FCV
infection when compared to a mock infection, and could reflect the fact that FCV infection
could induce the disassembly of pre-existing stress granules. However these results are difficult
to interpret as one could also argue that two mechanisms are overlapping: the inhibition of
152
stress assembly through an inhibition of stress signalling by FCV infection, and the active
dissolution of stress granules.
153
-Continued-
0hpi
2hpi
-SA/-FCV
+SA/-FCV
0.5 mM NaSAO2
+ FCV
p76 G3BP1 Overlay
p76 G3BP1 Overlay
p76 G3BP1 Overlay
p76 G3BP1 Overlay
p76 G3BP1 Overlay
+SA/+FCV
0.5 mM NaSAO2
+ FCV +SA/-FCV
0.5 mM NaSAO2
+ FCV +SA/+FCV
0.5 mM NaSAO2
+ FCV
154
6hpi
+SA/+FCV
0.5 mM NaSAO2
+ FCV
4hpi
Figure 3.24 Effect of FCV infection on pre-assembled stress granules. CRFK cells were
treated with 0.5 mM sodium arsenite (SA) for 1h, then infected with FCV at MOI of 0.2 for
several time points as noted on the panels. Cells were then immunostained for G3BP1
antibody (red) and p76 (green) as markers for stress granules and virus respectively,
followed by Alexa-555 and Alexa-488 secondary antibodies. Left panels show p76 alone
(green), middle panels show G3BP1 alone (red) and right panels show an overlay of both.
Cell nuclei were stained with ToPro-3 (blue). Cells were visualised using a ZEISS LSM
META confocal laser microscope. The bar indicates a size of 10 µm.
p76 G3BP-1 Overlay
p76 G3BP1 Overlay
p76 G3BP1 Overlay
p76 G3BP1 Overlay
+SA/-FCV
0.5 mM NaSAO2
+ FCV +SA/+FCV
0.5 mM NaSAO2
+ FCV +SA/-FCV
0.5 mM NaSAO2
+ FCV
155
F C V in fe c tio n (h p i)
Ce
lls
dis
pla
yin
g
str
es
s g
ran
ule
s (
%)
0 2 4 6 8
0
2 0
4 0
6 0
8 0
1 0 0
-S A /-F C V
-S A /+ F C V
+ S A /-F C V
+ S A /+ F C V
Figure 3.25 Quantification of the number of cells displaying stress granules upon sodium
arsenite (SA) treatment and FCV infection. CRFK were treated with 0.5 mM SA for 1h prior
infected with FCV at MOI of 0.2 for several time-points, stained for G3BP1 and p76 as
described in Figure 3.26. Confocal images were then used to quantify the numbers of cells with
stress granules using Image J software. Results shown are the percentage of cells displayed
SGs compared to the untreated uninfected, SA-treated and untreated infected, and are
representative of one replicate.
156
3.11 Effect of UV-inactivation on the inhibition of stress granule assembly
Because infection of CRFK cells with FCV led to the inhibition of SG formation
induced in both an eIF2 phosphorylation dependent and independent manner, this raised the
question whether this inhibition could be due to active viral replication or a cascade of cellular
events that would follow viral attachment.
To test these possibilities we used UV inactivation of FCV virus particles. Viral
particles were exposed to UV light (3 pulses of 4 min at 120,000 microjoules). To confirm the
inactivation, the ability of untreated and UV-inactivated viruses to replicate was then
determined using TCID50 assay. As shown in Figure 3.26, UV-inactivation strongly impaired
the ability of FCV to replicate with a reduction in titre from 1,28.10+6 to 0 TCID50/ml.
Subsequently untreated and UV-inactivated FCV particles were used to infect CRFK cells, and
6h post-infection these cells were challenged with sodium arsenite to induce SG assembly. The
formation of stress granules was monitored by immunofluorescence using G3BP1 as a marker
(Figure 3.27). As seen in Figure 3.27, UV-inactivation of FCV did not impair the induction of
SGs following sodium arsenite treatment, with 85.8% of cells displaying SGs as compared to
15.5% for untreated FCV particles (Figure 3.28). Therefore this strongly suggests that in order
to impair the assembly of stress granules, FCV particles need to enter the infected cells and
undergo active replication.
157
FC
V 6
h
UV
-in
act i
vate
d F
CV
6h
1 0 -1
1 0 0
1 0 1
1 0 2
1 0 3
1 0 4
1 0 5
1 0 6
1 0 7
TC
ID5
0/m
L
**
Figure 3.26 Effect of UV-inactivation on FCV replication. FCV were exposed to UV-light
as described in materials and methods, then incubated with CRFK cells for 6h. The viral titre
was determination using TCID50. Results shown are the titre of UV-inactivated virus compared
to the viral titre of active FCV, and are the mean of three independent experiments (+/-SE; **
is P ≤0.01).
158
-FCV/-SA
-FCV/+SA
+FCV 6hpi/
-SA
+FCV (UV) 6hpi/
-SA
+FCV (UV) 6hpi/
+SA
Figure 3.27 Effect of UV-inactivation on the inhibition of stress granules assembly. CRFK cells were infected with FCV or UV-inactivated FCV for 6h, treated with sodium arsenite (SA) to induce stress granule formation. Infected cells (C, D & E) and Mock cells (A & B) were then immunostained for G3BP1 (red) and p76 (green) as markers for stress granule and virus respectively, followed by Alexa-555 and Alexa-488 secondary. Left panels show p76 alone (green), middle panels show G3BP-1 alone (red) and right panels show an overlay of both. Cell nuclei were stained with ToPro-3 (blue). Cells were visualised using a ZEISS LSM META confocal laser microscope. The bar indicates a size of 10 µm.
A
B
C
D
E
overlay G3BP1 p76
overlay G3BP1 p76
overlay G3BP1 p76
overlay G3BP1 p76
overlay G3BP1 p76
159
-FC
V/-
SA
-FC
V/+
SA
+F
CV
(U
V)/
-SA
+F
CV
(U
V)/
+S
A
+F
CV
/+S
A
0
2 0
4 0
6 0
8 0
1 0 0
Ce
lls
dis
pla
yin
g
str
es
s g
ra
nu
les
(%
)
**
n s
Figure 3.28 Quantification of the number of cells displaying stress granules during
FCV or UV-inactivated FCV infection and sodium arsenite (SA) treatment. CRFK were
infected with FCV at MOI of 0.2 for 6h, stained for G3BP1 and p76 as described in Figure
3.22. Confocal images were then used to quantify the numbers of cells with stress granules
using Image J software. Results shown are the percentage of cells displaying SGs compared
to the control, and are the mean of three independent experiments (ns: not significant, ** is
P ≤0.01).
160
3.12 Discussion
Caliciviruses, in common with other RNA viruses, utilize the host translational
machinery in order to produce viral proteins. They however use a novel mechanism for
translation as they produce viral RNAs that lack a 5’ cap structure but instead have a virus-
encoded VPg protein covalently linked to the 5’ end of their mRNAs. VPg interacts with
eukaryotic initiation factors (eIF) and acts as a proteinaceous cap-substitute. Previously, we
have demonstrated that feline calicivirus (FCV) VPg interacts with eIF4E to direct translation,
whereas murine norovirus (MNV) VPg interacts with eIF4E and eIF4G, but only the interaction
with eIF4G is important for viral translation (Chaudhry et al., 2006; Chung et al., 2014).
Recently, results from our laboratory demonstrated that calicivirus replication is dependent on
the cell signalling pathways that control the phosphorylation of eIF4E, the MAPK pathway
(Royall et al., 2015). This also suggests that eIF4E phosphorylation is important in regulating
the translation of a subset of host mRNAs implicated in the antiviral response to calicivirus
infection (Royall et al., 2015). In addition to controlling transcription and the activity of the
protein synthesis machinery, recent studies have suggested that the assembly of stress granules
is central in orchestrating stress and antiviral responses to restrict viral replication (Onomoto
et al., 2014). Therefore, using MNV-1 and FCV as models, we investigated whether
caliciviruses control the host response to infection by modulating stress granule responses.
161
3.12.1 Absence of stress granules in stressed and MNV-infected RAW264.7 or BV-2
macrophages.
Using different MNV-permissive cells, including RAW264.7 and BV-2 cells, we could
not detect the assembly of SGs during MNV1 infection (Figure 3.7, 3.14-3.16) and only limited
assembly of granules containing G3BP1, but not TIA-1, and eIF3η, following oxidative stress
with sodium arsenite. These results may reflect a deficiency in RAW264.7 and BV-2 cells,
both of which are macrophages, in their ability to form stress granules. Alternatively, this could
reflect that the antibodies used could be species-specific and not cross-react with other species
such as murine.
It has been shown previously that the overexpression of a protein regulating the
inflammatory response and immune homeostasis, MCPIP1 (monocyte chemotactic protein-
induced protein 1), can impair the assembly of SGs in response to cellular stresses. Whilst
overexpression of MCPIP1 sensitized RAW264.7 cells to stress-induced apoptosis, MCPIP1-
deficient cells are resistant to stress-mediated apoptosis (Qi et al., 2011). MCPIP1 functions
as a deubiquitinase (DUB) to inhibit several signalling pathways such as c-Jun N-terminal
kinase (JNK) signalling pathways (Liang et al., 2010). In addition, MCPIP1 displays RNase
activity towards some inflammatory cytokine mRNAs and viral RNA (Lin et al., 2013;
Mizgalska et al., 2009). As a consequence MCPIP1 inhibits replication of HCV, JEV and
DENV (Lin et al., 2013). However, the ability of MCPIP1 to inhibit SG formation is dependent
on its deubiquitinating enzyme activity and it can impair the assembly of SGs following
oxidative stress with sodium arsenite (Qi et al., 2011). It was suggested that stress results in
the overexpression of MCPIP1 which in turn inhibited arsenite-induced eIF2α phosphorylation.
Therefore, we could hypothesize that the absence of SGs during MNV-1 infection in
this cell type reflects a mechanism in which MNV-1 infection blocks SG formation through
162
the overexpression of MCPIP1, and this could be conserved in BV-2 cells. To fully understand
the role of MNV-1 in SG assembly future work should include the identification of a cell-line
supporting MNV-1 infection and being responsive to oxidative stress, so that positive control
induction experiments are allowed. Alternatively, an MCPIP1-deficient RAW264.7 cell line
could be used for MNV infection and subsequent detection of stress granules, in combination
with monitoring the levels of MCPIP1 by immunoblotting during MNV infection.
3.12.2 FCV replication impairs stress granule assembly during infection
The activation of stress activated kinases by viral infection, mainly PKR, but also
PERK, results in the phosphorylation of eIF2α (Holcik & Sonenberg, 2005). This in turn
prevents the recycling of the ternary complex tRNAiMet-GTP-eIF2, thereby stalling the
initiation of translation resulting in a shutdown of protein synthesis and causing SGs to
accumulate in the cytoplasm (Anderson & Kedersha, 2008). Using G3BP1 and PABP as
markers for SGs, and CRFK and FEA as cells supporting FCV replication, we could not find
evidence of stress granule induction during FCV infection (Figure 3.8). Only uninfected cells
displayed SGs and this observation is dissected further in Chapter 4.
The resulting stoppage in protein synthesis following SG assembly is problematic for
viruses as all viruses rely on the host cell protein synthesis machinery for production of their
proteins. Consequently viruses have evolved many strategies to ensure that viral mRNA
translation can continue by disrupting SG formation during infection or exploiting SGs for their
replication (Montero & Trujillo-Alonso, 2011; Reineke & Lloyd, 2013; Valiente-Echeverria et
al., 2012). The absence of SGs in infected cells during FCV infection could therefore reflect
an inhibition of stress responses during FCV infection.
163
Amongst the different stresses that lead to translational arrest and the formation of SGs,
the most common is dependent on phosphorylation of the α-subunit of eukaryotic initiation
factor 2 (eIF2) at Ser51(Kedersha et al., 1999; McEwen et al., 2005). Typically, sodium
arsenite activates the HRI eIF2α kinase causing oxidative stress-induced phosphorylation of
eIF2α (McEwen et al., 2005). However, translational arrest can also occur independently from
eIF2α phosphorylation through the disruption of the eIF4F complex (Dang et al., 2006; Emara
et al., 2012). Pateamine A (PatA) inhibits eukaryotic translation initiation eIF4A RNA helicase
activity while hydrogen peroxide induces 4EBP hypo-phosphorylation and sequestration of
eIF4E from eIF4F (Emara et al., 2012).
Using sodium arsenite and hydrogen peroxide as stress inducers, representative of the
eIF2α-dependent and independent pathways, respectively, my results showed that FCV
infection impairs SG assembly (Figure 3.17 and 3.18). Only 7% of FCV-infected cells
challenged with sodium arsenite displayed SGs, compared with 83.4% for sodium arsenite-
stressed cells. Likewise 7.7% of FCV-infected cells challenged with hydrogen peroxide
displayed SGs, compared with 87.7% for hydrogen peroxide-stressed cells. Therefore this
would suggest that FCV infection impairs the assembly of stress granules triggered both in an
eIF2α dependent and independent manner. However, because some studies suggested that
H2O2 does not induce SGs (Arimoto et al., 2008; Kedersha & Anderson, 2007), additional
proof of FCV inhibition of eIF2-independent SG assembly could be provided using
Pateamine A or Hippuristanol as inhibitors of eIF4A activity.
The strategies developed by viruses to inhibit stress granules include: (i) proteolytic
degradation of RNA granule factors by viral proteases, (ii) regulation of eIF2α phosphorylation
and (iii) co-opting of RNA granule factors for a new role in viral replication or sequestration
reviewed in (Reineke & Lloyd, 2013). Because FCV infection seems to inhibit both eIF2-
dependent and independent pathways, and given that a previous Ph.D student at Surrey reported
164
eIF2α phosphorylation during FCV infection (Alessandro Natoni, thesis manuscript), it is
likely that SG assembly is inhibited by either (i) proteolytic degradation of RNA granule factors
by viral proteases and (iii) co-opting of RNA granule factors for a new role in viral replication
or sequestration. This would exclude an inhibition mechanism whereby eIF2α phosphorylation
is regulated during infection. These two mechanisms rely on viral products, RNA or proteins
to actively impair SG assembly. Because infection with UV-inactivated virus leads to efficient
SG aggregation following sodium arsenite treatment, this indicates that FCV replication, and
therefore either viral protein(s) or RNA is required to mediate SG inhibition (Figure 3.27).
Several previously identified inhibition mechanisms involve the targeting of the SG-
nucleating protein G3BP1. Non-structural proteins of several alphaviruses interact with G3BP1
in viral complexes that contain nsp3 or nsp2 and nsp4 (Cristea et al., 2010; Fros et al., 2012).
SFV hijacks G3BP1 from SGs and relocates it into viral replication complexes via an
interaction with the C-terminal repeat domains of nsp3, disassembling SGs (Panas et al., 2015;
Panas et al., 2012). As a consequence, SG induction by the eIF2-independent inducer
Pateamine A is blocked (Panas et al., 2012). Similarly, SGs do not form upon infection with
JEV and the JEV core protein directly binds Caprin1 preventing mRNA recognition and SG
formation (Katoh et al., 2013). Viral proteinase from picornaviruses can disperse stress
granules through the cleavage of the SG-nucleating protein G3BP1 by the conserved 3C
proteinases (3Cpro) (Ng et al., 2013; White et al., 2007). Caliciviruses encode a proteinase
designated as a 3C-like proteinase (3CLpro) because of its sequence similarity to the
picornavirus 3Cpro in the active site region (Sosnovtseva et al., 1999), so it is tempting to
speculate that the FCV inhibition of SGs might results from G3BP cleavage. However, using
cell lysates from FCV-infected cells that I generated at the end of my laboratory work,
preliminary immunoblotting experiments suggest that G3BP1 is not cleaved during infection.
However, we cannot exclude that the 3CLpro plays a role in SG disruption by cleaving another
165
SG component. To explore further the role of the FCV 3CLpro, further studies should take
advantage of the possibility of overexpressing a wild-type FCV 3CLpro or an inactive mutant
in CRFK cells (Ian Goodfellow, personal communication). Examination of SG assembly
following sodium arsenite treatment under both conditions should reveal whether the FCV
3CLpro plays any role in the inhibition of SG observed during FCV infection.
Another effective way by which viruses can inhibit stress granule assembly is by
redirecting key SG nucleating proteins into various viral replication foci or other subcellular
localization. For example, IAV can inhibit SG formation by several mechanisms which include
the inactivation of PKR through its binding with non-structural protein 1 (NS1), which in turn
prevents eIF2α phosphorylation and blocks SG formation, while another mechanism requires
the nucleoprotein (NP) without affecting eIF2a phosphorylation (Khaperskyy et al., 2014). In
addition, the host-shutoff protein, namely the polymerase-acidic protein-X (PA-X), can
strongly inhibit SG formation by inducing the concomitant depletion of cytoplasmic poly (A)+
mRNA and the accumulation of poly (A)-binding protein in the nucleus, which impairs the
assembly of PABP-containing SGs later in infection despite eIF2α phosphorylation
(Khaperskyy et al., 2014). Using PABP as SG marker, it was also evidenced that FCV-infection
impairs the assembly of SGs following oxidative stress (Figure 3.10).
Several studies have previously reported that nucleolin, a ubiquitous multifunctional
nucleolar shuttling phosphoprotein, can interact with proteins that are known components of
SGs such as HuR, Staufen, PABP and G3BP1(Isabelle et al., 2012; Tominaga et al., 2011;
Villace et al., 2004; Zheng et al., 2008). Furthermore, nucleolin interacts with the 3’UTR of
NoV and FCV genomic RNA, and the viral RdRp, participating in the replication complex
localized in perinuclear locations (Cancio-Lonches et al., 2011). Therefore, another hypothesis
could be that during infection FCV destabilizes SG assembly by relocalizing SG components
166
to replication complexes. Further studies should aim at dissecting the potential co-localisation
of SG markers within FCV replication complexes using immunofluorescence studies.
Overall the results obtained in this chapter suggest that FCV infection has the ability to
prevent stress responses and avoid the inhibition of protein synthesis, thus enhancing virus
replication. Further studies should be performed to further confirm the impact of FCV infection
on the activation of various virus-related stress pathways previously linked to SGs such as
members of the PKR, RIG-I or c-JUN pathways using immunoblotting.
167
Chapter 4
Paracrine induction of Stress
Granules by
FCV-infected cells
168
4.1 Generation of virus-free cell culture supernatant.
The results from the previous chapter strongly suggest that FCV infection impaired
the ability of CRFK cells to assemble SGs during infection. However, during the infection
SGs could be detected in cells that displayed no sign of infection (revealed by p76 staining)
but resided in the vicinity of infected cells, with 15% of cells displaying SGs at 4h post-
infection. This may reflect a cell-to-cell, paracrine, induction via a chemical or biological
signal of SG formation to dampen viral propagation and/or mount an antiviral response in
uninfected cells.
The only studies that have revealed a mechanism that may be related to this originate
from the observation of mosquito cells infected with Sindbis virus or Semliki Forest virus
(Newton & Dalgarno, 1983; Riedel & Brown, 1979). Cells from the mosquito species Aedes
albopictus, persistently infected with Sindbis virus, produce a low molecular-weight
biomolecule with antiviral activity, capable of specifically affecting Sindbis virus replication.
Unlike interferon, this antiviral agent is virus-specific and cell-specific. Furthermore its
activity is inactivated by treatment with protease K enzyme and heat, but not in response to
anti-Sindbis antibodies (Riedel & Brown, 1979). A similar effect on virus production was
also observed in mosquito cells treated with antiviral material released into the medium from
Semliki Forest virus (SFV)-infected cells. Again this antiviral medium was inactivated by
heat and proteinase K and is specifically associated with alphaviruses and has no significant
effect on other types of arbovirus (Newton & Dalgarno, 1983). However, despite being
identified more than 30 years ago, no mechanistic details have been further elucidated.
169
Therefore, we reasoned that FCV-infected cells may be releasing a signal that would
be able to act as a paracrine inducer of SG formation. To test this hypothesis, we first
generated virus-free supernatant from FCV-infected cells. To this end, CRFK cells were
mock or FCV infected for 6h. The cell culture supernatant was then collected and the viral
particles were UV-inactivated, precipitated using PEG3350 and NaCl and eliminated by
centrifugation. Subsequently, the collected supernatant was assayed for the presence of viral
particles by measuring viral titres using TCID50 assays, and incubating CRFK cells with
supernatants followed by virus detection using immunofluorescence. As shown in Figure 4.1,
viral replication could not be detected by TCID50 following virus elimination as described
above. Thus, we assumed that FCV particles were eliminated from the cell culture
supernatants.
170
FC
V 6
h
FC
V-
su
pern
ata
nt
6h
1 0 0
1 0 1
1 0 2
1 0 3
1 0 4
1 0 5
1 0 6
1 0 7
1 0 8
1 0 9T
CID
50
/mL
Figure 4.1 Detection of FCV replication by TCID50 assay following virus elimination.
CRFK cells were incubated with FCV-infected cell supernatant for 6h, viral titre was then
determined by TCID50 assay. Results show viral titre from supernatant-treated cells compared
to the viral titre from untreated infected cells.
171
4.2 Effect of FCV-infected cell supernatant on Stress granule formation.
Having confirmed that we were able to collect cell culture supernatants free of viral
particles, we proceeded to assay whether these were able to mediate SG formation in
uninfected CRFK cells. To this end, CRFK cells were mock- or FCV-infected for 6h and the
supernatant collected. Viral particles were then eliminated using the procedure described in
section 2.16 (Materials and Methods). The supernatants were then incubated with uninfected
CRFK cells for 1 and 6h. Subsequently the presence of SGs was monitored using
immunofluorescence with G3BP1 as marker, and p76 as marker for viral replication (as a
negative control).
As shown in Figure 4.2, the incubation of CRFK cells with cell supernatant collected
from mock-infected cells did not induce the formation of SGs, reflected by the absence
G3BP1 cytoplasmic foci, either following 1h or 6h of incubation (Figure 4.2, Figure 4.3). In
contrast, the supernatant from FCV-infected cells, was able to induce SG formation in CRFK
cells, with 26.2% of cells displaying SGs after 1h of incubation and 41.7% after 6h.
Furthermore, immunofluorescence microscopy confirmed that no viral particles were present
as reflected by the absence of signal when staining for the viral marker p76 (Figure 4.2).
Therefore these results suggest that FCV-infected cells are able to mediate the release of a
signal that induces SG assembly in neighbouring cells.
172
Figure 4.2 Induction of stress granules by cell supernatant. CRFK cells were incubated with
supernatant from mock or FCV-infected cells for 1h and 6h and then immunostained for G3BP1
antibody (red) and p76 antibody (green) as markers for stress granule and virus respectively,
followed by Alexa-555 and Alexa-488 secondary. Left panels show p76 alone (green), middle
panels show G3BP1 alone (red) and right panels show an overlay of both. Cell nuclei were
stained with ToPro-3 (blue). Cells were visualised using a ZEISS LSM META confocal laser
microscope. The bar indicates a size of 10 µm.
Overlay G3BP1 p76
Overlay G3BP1 p76
Overlay G3BP1 p76
Overlay G3BP1 p76
Overlay G3BP1 p76
- sup
+ mock-sup (1h)
+ FCV-sup (1h)
+ mock-sup (6h)
+ FCV-sup (6h)
173
mock-s
up 1
h
FC
V-s
up 1
h
mock-s
up 6
h
FC
V-s
up 6
h
0
2 0
4 0
6 0
8 0
1 0 0
Ce
lls
dis
pla
yin
g
str
es
s g
ran
ule
s (
%)
***
*
Figure 4.3 Quantification of the number of cells displaying stress granules following
treatment with cell supernatant. CRFK cells were incubated with mock or FCV-infected
cell supernatants (sup) and immunostained for G3BP1 as described in Figure 4.1. Image J
software was used to quantify the number of cells displaying SGs. Results show the
percentage of cells with SGs induced by supernatant from FCV-infected cells compared to
either the mock-infected supernatant or untreated cells, and are the mean of three independent
experiments (+/-SE; *** is P≤0.001 for 1h and * is P ≤0.5 for 6h).
174
4.3 Effect of interferon α on stress granule formation in CRFK cells
The production of interferon can be induced in animal cells as the innate immune
response by a variety of antigens and especially by virus infection (Friedman, 1977; Randall
& Goodbourn, 2008). Viral RNA activates signalling pathways that result in the production
of interferons (IFNs) and IFN-stimulated genes which in turn trigger an intracellular antiviral
state that leads to inhibition of a wide range of viruses (Onomoto et al., 2014; Umbach &
Cullen, 2009). Recognition of RNA viruses by TLR7/8 results in the induction of IFN
production via the activation of IRF5 and IRF7 signalling cascades, which substantially leads
to the activation of IFN stimulated genes that mediate the antiviral response (Perry et al.,
2005). Previously it was shown that addition of type I IFN (IFNβ) to cells infected with
measles virus enhanced the assembly of SGs, and IFNβ treatment alone, in the absence of
infection, induced SG formation in ADAR1-deficient cells(John & Samuel, 2014). Moreover
it was reported that IFNβ treatment of HCV-infected cells strongly enhances HCV-induced
phosphorylation of PKR and eIF2α, resulting in increasing SG formation (Garaigorta et al.,
2012; Ruggieri et al., 2012). Based on the general antiviral properties of interferon and these
studies, we reasoned that FCV-infected cells may simply release IFN as a response to
infection, and that this could act as a potent paracrine inducer of SG assembly.
To test this hypothesis, we used commercial feline IFN to investigate the effect of
type I IFN on the accumulation of SGs in CRFK cells. Previous studies have shown that
CRFK cells are responsive to IFN treatment (Baldwin et al., 2004; Harun et al., 2013). CRFK
cells were treated with increasing amount of recombinant feline IFN from 150 IU/ml to
3000 IU/ml and the formation of SGs was monitored by immunofluorescence using G3BP1
as SG marker. As shown in Figure 4.4 and 4.5, the addition of increasing amount of IFNdid
175
not induce any significant increase in the number of SGs, as reflected by the absence of
cytoplasmic G3BP1 foci (Figure 4.4). Therefore, we speculate that type I IFN, at least IFN,
is unlikely to play a role in the paracrine induction of SGs by FCV-infected cells. We
however acknowledge that we did not test directly whether the recombinant IFN used
triggered a response in CRFK cells, however this was published before as mentioned above.
176
0 IU/ml
150 IU/ml
200 IU/ml
300 IU/ml
3000 IU/ml
overlay G3BP1 ToPro-3
overlay G3BP1 ToPro-3
overlay G3BP1 ToPro-3
overlay G3BP1 ToPro-3
overlay G3BP1 ToPro-3
Figure 4.4 Effect of interferon treatment on stress granule formation. CRFK cells were
incubated with increasing concentrations of IFN-α for 6h, then immunostained for G3BP-1
antibody as marker for stress granule. Cell nuclei were stained with ToPro-3 (blue) (left panels)
and middle panels show G3BP alone (red) and right panels show an overlay of both. Cells were
visualised using a ZEISS LSM META confocal laser microscope. The bar indicates a size of 10
µm.
177
0150
200
300
3000
0
1 0
2 0
3 0
4 0
5 0
IF N ( IU /m L )
Ce
lls
dis
pla
yin
g
str
es
s g
ran
ule
s (
%)
n s
n s
n s
n s
Figure 4.5 Quantification of the number of cells displaying stress granules following
treatment with IFNα. CRFK cells were incubated with increasing concentrations of IFNα and
immunostained for G3BP1 as described in Figure 4.4. Image J software was used to quantify
the number of cells displaying SGs. Results show the percentage of IFNα-treated cells with SGs
compared to mock-treated cells.
178
4.4 Effect of Ribonuclease A (RNase A) on the ability of FCV-infected cell supernatant
to induce stress granule assembly
The accumulation of an RNA replication intermediate during infection leads to the
activation of PKR, which then triggers several downstream effectors involved in the
production of pro-inflammatory cytokines and stress pathways resulting in SG assembly
(Bonnet et al., 2000; Onomoto et al., 2014). Furthermore, in response to infection, host and
viral miRNAs play an important role in regulating expression of viral and host genes
involved in the antiviral response (Emde & Hornstein, 2014; Leung & Sharp, 2010).
Therefore RNA-derived molecules can play key roles in antiviral signalling.
Given that type I IFN is not responsible for the induction of SGs mediated by FCV-
infected cell supernatant, we hypothesise that this as yet unidentified messenger could have
an RNA origin, as various forms of RNA can act as signals for antiviral response. To test this,
we treated the supernatant with RNase A as described in the methods section, a pyrimidine-
specific endonuclease which is commonly used to digest RNA polymers. The ability of the
RNase A-treated supernatant to induce SG assembly in CRFK cells was then monitored using
immunofluorescence microscopy using G3BP1 as SG marker (Figure 4.6). As shown in
Figure 4.6, the digestion of RNA polymers had no effect on the ability of FCV-infected cell
supernatant to induce SG assembly with 25.3% of cells displaying SGs compared with 31%
for untreated supernatant (Figure 4.6, Figure 4.7). Therefore these results suggest that free
RNA polymers are not involved in the signalling from infected to uninfected cells that
triggers SG assembly.
179
Overlay G3BP1 ToPro-3
Overlay G3BP1 ToPro-3
Overlay G3BP1 ToPro-3
Untreated cells
+ FCV-sup/
- RNAse A
+ FCV-sup/
+ RNAse A
Figure 4.6 Effect of RNase A treatment on stress granule formation. FCV-infected cell
supernatants were treated with 7U of RNAse A, and incubated with CRFK cells for 6h, then
immunostained for G3BP1 antibody as marker for stress granule (red). Cell nuclei were stained
with ToPro-3 (blue) (left panels) and middle panels show G3BP1 alone (red) and right panels
show an overlay of both. Cells were visualised using a ZEISS LSM META confocal laser
microscope. The bar indicates a size of 10 µm.
180
FC
V-s
up
FC
V-s
up +
RN
Ase A
Untr
eate
d c
ells
0
1 0
2 0
3 0
4 0
5 0C
ell
s d
isp
lay
ing
str
es
s g
ran
ule
s (
%)
n s
Figure 4.7 Quantification of the number of cells displaying stress granules following
treatment with RNase A. CRFK cells were incubated with 7U RNAse A and
immunostained for G3BP1 as described in Figure 4.6. Image J software was used to
quantify the number of cells displaying SGs. Results show the percentage of cells with
SGs induced by supernatant (sup) from FCV-infected cells compared with cells treated
with FCV-cell supernatant plus RNase, and are the means of three independent
experiments (+/-SE; is ns: not significant).
181
4.5 Effect of Heat-Shock on the ability of FCV-infected cell supernatant to induce stress
granule assembly
The infection of cells by viruses is a trigger for a large array of innate immune
responses. Several immune receptors expressed on the surface of cells or internally can ignite
that immune response, the major one being Toll-like receptors (TLRs), retinoic-acid
inducible gene I (RIG-I) and NOD-like receptors (NLRs). RNA viruses can activate the TLRs
displayed on endosomal compartments such as TLR3, TLR7, TLR8 and TLR9 (Pichlmair &
Reis e Sousa, 2007), while replication intermediates also activate the PKR pathway (Garcia et
al., 2006). These events result in the production of many pro-inflammatory cytokines and
chemokines (such as TNF, IL-1 or IL-8) and IFN that are secreted and can amplify the
immune response via both autocrine and paracrine effects, and further impact on the outcome
of infection (Rouse & Sehrawat, 2010).
Therefore we investigated whether the induction of SGs by virus-free supernatants
could be mediated by the release of protein messengers, putatively an immune mediator of a
proteinaceous nature. To test this hypothesis, we performed heat-shock of the supernatant to
denature proteins by heating supernatants at 65°C for 10 or 20 min. The ability of the heat-
shocked supernatant to induce SG assembly in CRFK cells was then monitored using
immunofluorescence and G3BP1 as SG marker (Figure 4.8). As shown in Figure 4.8, the
denaturation of proteins impaired the ability of FCV-infected cell supernatant to induce the
formation of SGs. This was reflected by a reduction in G3BP1 cytoplasmic foci following 10
min inactivation, dropping from 30.9% to 0.5%, and a complete disappearance of SGs
following 20 min of inactivation (Figure 4.8, Figure 4.9). Therefore these results could
182
suggest that the signalling from FCV-infected to uninfected cells involves a messenger of
proteinaceous nature.
183
Untreated cells
FCV-sup
FCV-sup + 10 min heat
shock
FCV-sup + 20 min heat
shock
Figure 4.8 Effect of heat shock on stress granule formation. FCV-infected cell
supernatants (FCV-sup) were exposed to heat shock for 10 or 20 min, incubated with
CRFK cells for 6h, then immunostained for G3BP1 antibody as markers for stress
granule (red) followed by Alexa-555 secondary antibody. Cell nuclei were stained
with ToPro-3 (blue) (left panels) and middle panels show G3BP alone (red) and right
panels show an overlay of both. Cells were visualised using a ZEISS LSM META
confocal laser microscope. The bar indicates a size of 10 µm.
Overlay G3BP1 ToPro-3
Overlay G3BP1 ToPro-3
Overlay G3BP1 ToPro-3
Overlay G3BP1 ToPro-3
184
FC
V-s
up
FC
V-s
up
+ H
eat-
sh
ock 1
0 m
in
FC
V-
su
p+ H
eat-
sh
ock 2
0 m
in
Un
treate
d c
ells
0
2 0
4 0
6 0
8 0
1 0 0C
ell
s d
isp
lay
ing
str
es
s g
ra
nu
les
(%
)
* * *
* * *
n s
Figure 4.9 Quantification of the number of cells displaying stress granules following
treatment with heat-shocked supernatant. Confocal microscopy was used to detect the
effect of heat shock on stress granule formation as described in Figure 4.8, Image J software
was then used to quantify the number of cells displaying SGs following treatment with heat-
shocked supernatant. Results show the percentage of cells with SGs induced by heat-shocked
FCV-infected cell supernatant compared with cells treated with unheated FCV-infected cell
supernatant, and are the means of three independent experiments (+/-SE; *** is P ≤0.001, ns
not significant).
185
4.6 Effect of FCV-infected cell supernatant on a wide range of cells’ ability to assemble
stress granules
Having now proposed that FCV-infected CRFK cells release a proteinaceous
messenger that can mediate the assembly of SGs, we next ask whether this messenger could
have a ubiquitous action and induce the assembly of SGs in other cell lines. To test this
hypothesis, we incubated the virus-free supernatant of FCV-infected CRFK cells with
different cells, including FEA (feline cells supporting FCV replication), 293T (a human
embryonic kidney cell line supporting FCV replication), RAW264.7 (murine macrophage
cells supporting MNV-1 replication), BV-2 (murine microglia cells supporting MNV-1
replication) and J774 (murine macrophage cells supporting MNV-1 replication) and THP-1
(human macrophage cells), for 6h. The presence of SGs was then monitored by
immunofluorescence using G3BP1 as marker. G3BP1 cytoplasmic foci, representative of
SGs, were clearly detected in FEA and 293T cells with 71.6% and 57.3% of cells displaying
SGs, respectively (Figure 10, Figure 11). In stark contrast, no SG induction could be detected
in RAW264.7 and BV-2 cells (Figure 10, Figure 11) as well as THP-1 and J774 (data not
shown). Of note these cell lines are all murine or human macrophage cell lines. Therefore,
although the supernatant of FCV-infected CRFK cells has the ability to induce SG formation
in other cell types, it does seem to exhibit a level of cell specificity, and could be related to
the fact that these cells support FCV replication.
186
Figure 4.10 Effect of FCV-infected cell supernatant on a wide range of cells. 293T, FEA, RAW264.7, BV-2,
THP-1 and J774 cells were grown as described in Materials and Methods, then treated with supernatant from
FCV-infected cells (FCV-sup) for 6h. Untreated and treated cells were then immunostained for G3BP1 (red) and
p76 (green) as markers for stress granule and virus respectively, followed by Alexa-555 and Alexa-488
secondary antibodies. Left panels show untreated cells (-FCV-sup), right panels show treated cells (+FCV-sup).
Cell nuclei were stained with ToPro-3 (blue). Cells were visualised using a ZEISS LSM META confocal laser
microscope. The bar indicates a size of 10 µm.
G3BP1 G3BP1
G3BP1 G3BP1
THP-1
J774
G3BP1 G3BP1
293T
- FCV-sup + FCV-sup
G3BP1
FEA
G3BP1
G3BP1 G3BP1
RAW264.7
BV-2
G3BP1 G3BP1
187
293 c
ells
FE
A c
ells
RA
W 2
64.7
cells
BV
-2 c
ells
TH
P-1
cells
J774 c
ells
0
2 0
4 0
6 0
8 0
1 0 0
Ce
lls
dis
pla
yin
g
str
es
s g
ra
nu
les
(%
)
U n tre a te d c e lls
F C V -s u p
Figure 4.11 Quantification of the number of cells displaying stress granules following
treatment of a wide range of cells with FCV-infected cell supernatant. CRFK cells
were incubated with mock- or FCV-infected cell supernatants and immunostained for
G3BP1 as described in Figure 4.10. Image J software was used to quantify the number of
cells displaying SGs. Results show the percentage of cells with SGs induced by
supernatant from FCV-infected cells compared with untreated cells.
188
4.7 Discussion
4.7.1 FCV infection leads to paracrine induction of SG assembly
Because of the ability of viruses to trigger the formation of SGs and cause
translational shut-off, it has been proposed that SGs could exert a specific antiviral effect,
both on viral mRNA and/or by regulating mRNA contributing to the antiviral response
(Onomoto et al., 2014). Following our observation that SGs can be detected in some
uninfected cells during FCV infection, we hypothesize that FCV-infected cells may release a
signalling molecule that could trigger SG assembly in neighbouring cells, which would be
consistent with this proposed antiviral role for SGs.
To test this hypothesis, I generated virus-free supernatants from mock- and FCV-
infected cells. These supernatants were first tested to confirm the absence of viral replication
by determining viral titre using a TCID50 assay (Figure 4.1) as well as an
immunofluorescence assay to detect viral protein (Figure 4.2). Neither method detected viral
protein or viral replication confirming that the supernatants were indeed virus-free.
The supernatants were then added to CRFK cells and the presence of SGs was
examined by immunofluorescence microscopy (Figure 4.2). While the supernatant from
mock-infected cells did not trigger SG formation, 26% of cells treated with virus-free
supernatant from FCV-infected cells formed SGs after 1h and 42% after 6h (Figure 4.3).
These results suggest that FCV-infected cells release a soluble mediator that induces SG
assembly in neighbouring cells. This paracrine induction of SGs in uninfected cells has never
been described and may reflect a novel regulatory mechanism to control stress during
infection by modulating mRNA metabolism.
189
4.7.2 Different types of stress granules can be assembled in response to infection
Recent studies have suggested that SGs have an antiviral role and function as a
cellular platform to initiate innate immune responses (Onomoto et al., 2014). SGs can serve
as sentinel mechanisms for IFN activation by promoting the condensation of IFN signalling
moieties with their viral RNA ligands (Khaperskyy et al., 2014; Langereis et al., 2013;
Onomoto et al., 2012; Yoo et al., 2014). In addition to canonical SG markers, these antiviral
SGs contain viral RNA and nucleoproteins. Thus, stress granules may be important in
priming neighbouring cells against infection by (i) suppressing viral replication through
inhibition of viral protein synthesis, and (ii) serving as a platform to facilitate IFN production.
In addition to these, viruses have also been linked to the formation of antiviral granules
(AVGs) (Rozelle et al., 2014). The disruption of Poxvirus replication triggers the formation
of host-protein dense antiviral granules (AVGs) which differ from SGs by their localization
to cytoplasmic viral factories and by their composition (Rozelle et al., 2014). While these
AVGs share many components with SGs, they are characterised by the absence of eIF3 and
40S subunits, and their stability in the presence of translation elongation inhibitors. This
suggested that AVGs may not act as sorting sites for mRNAs but exists to restrict viral
translation. Overall a link between antiviral immunity and stress granules is clear, there seem
to be different specific responses linked to different types of RNA granules.
To investigate the nature of the stress granules induced by virus-free supernatant,
further studies should aim at investigating whether these are dissolved following
cycloheximide treatment, as canonical SGs. Then, the composition of these SGs should be
characterized using immunofluorescence microscopy to detect a variety of canonical SG
markers (G3BP1, Caprin, TIA1, PABP, eIF4G), IFN signalling factors previously found to
accumulate in avSGs (RIG-I, MDA5, PKR) and markers that accumulate in SGs but not
AVGs (40S, eIF3 and poly(A)+ mRNA using digoxigenin-labeled oligo-d(T) and anti-
190
digoxigenin antibody). Furthermore, because canonical SG assembly mostly results from
translational arrest, further work should aim at investigating whether the assembly of SGs
mediated by the supernatant of infected cells is the consequence of a translational shut-off.
To determine whether translation is inhibited in the subset of cells that form SGs,
visualization of de novo protein synthesis by metabolic incorporation of the amino acid
analog L-azidohomoalanine (AHA) could be employed. After cell fixation, proteins that had
incorporated AHA can be conjugated with a fluorescent probe, such as Alexa Fluor 594,
using a copper-catalysed reaction and labelling any nascent peptide with AHA-594 as
described previously (Rozelle et al., 2014). As a control, cells should be treated with the
translation inhibitor cycloheximide, which should completely block AHA-594 staining. Cells
would also be stained for stress granule markers, such as G3BP1, to score both stress granule
formation and active translation in cells. This would identify any link between inactive
translation and SG induction, providing evidence as to whether virus-free supernatant
mediates a translational shut-off.
4.7.3 Could this paracrine induction of SGs reflect an antiviral mechanism?
Previously, some studies have described the release of a low molecular weight
antiviral agent into the medium of A.albopictus cells persistently infected with Sindbis virus
(Riedel & Brown, 1979), or Semliki Forest virus (Newton & Dalgarno, 1983). This agent was
able to reduce the normal viral yield. Treatment of mosquito cultures with the supernatant
containing the antiviral agent before infection prevents the normal production of high virus
yield (Newton & Dalgarno, 1983; Riedel & Brown, 1979). However, the antiviral activity
was sensitive to proteinase K and heat, being rapidly inactivated at 56˚C. These studies
reported that the antiviral agent was also cell and virus specific. While this antiviral activity
191
has been shown to selectively affect Sindbis virus production in mosquito cells, it had no
effect on virus production in BHK-21 cells (Condreay et al., 1988; Condreay & Brown, 1988;
Riedel & Brown, 1979). In addition, Kanthong et al., 2010 discovered that cell-free
supernatant solutions from C6/36 mosquito cell cultures are capable of destabilizing
persistently-infected cultures with Dengue virus. The authors proposed that the cell-free
supernatant contains small polypeptides with cytokine-like activity resulting in an effect on
virus yield and to induce a protective response against DENV-2 virus infection in naïve cells
(Kanthong et al., 2010).
In agreement with these data, our results report that the ability of virus-free
supernatant to induce SGs was sensitive to heat treatment at 65˚C (Figure 4.8). In addition,
our results demonstrated that the effect of this supernatant was not due to RNA, as RNase
treatment had no impact (Figure 4.7). A subsequent experiment was carried out to determine
the nature of this antiviral agent contained within the supernatant. The fact that virus infected
cells could induce an antiviral state in surrounding host cells during most viral infections, led
us to the hypothesis that the induction of SGs by supernatant may be due to interferon
activity. However, treating CRFK cells with feline IFN did not induce SG assembly, showing
that release of IFN from infected cells is unlikely to be responsible for the paracrine SG
induction (Figure 4.4).
To further investigate whether the assembly of SGs in uninfected cells reflects the
induction of a protective antiviral effect, CRFK cells could be treated with virus-free
supernatant from mock or FCV-infected cells and then challenged with FCV. Viral
replication would then be quantified by measuring viral titre using TCID50 assay. The
antiviral state of the cells would then be characterized further by measuring IFN levels using
qRT-PCR. In addition the conservation of this mechanism could be investigated by analysing
192
the ability of virus-free supernatant from cells infected with other viruses to induce SGs,
using model arboviruses and herpesviruses studied at Surrey.
4.7.4 Investigating the nature of the soluble mediator(s) triggering stress granule
assembly
To date, the only example of a paracrine inducer of SG assembly is angiogenin, an
angiogenic factor released by tumour cells (Yamasaki et al., 2009). Angiogenin is a secreted
stress-activated ribonuclease producing tRNA-derived stress-induced RNA (tiRNAs) to
promote translational repression and induce SG assembly (Emara et al., 2010). To investigate
whether angiogenin plays a role in the paracrine induction of SGs in bystander cells, extracts
prepared from CRFK cells treated with virus-free supernatants of mock-infected or FCV-
infected cells could be separated on a denaturing gel and analyzed to visualize stress-induced
small RNA by northern blotting with cDNA probes complementary to the 5’ and 3’ end of
tRNA. In addition, CRFK cells could be transfected with control or Dharmacon SMART pool
siRNA targeting angiogenin to reduce its expression before investigating the effect on virus-
free supernatant ability to induce SGs.
We cannot however rule out that the paracrine inducer of SG assembly could have a
different nature: nucleic acid, such as RNA fragments of viral origin, cellular miRNAs, other
proteins such as cytokines or signalling proteins, or more complex structures such as
exosome particles which can deliver antiviral sensors to uninfected cells (Kalamvoki et al.,
2014). It has been proposed that exosomes may be responsible for transferring anti-viral
signal between cells (Li et al., 2013; Schorey & Bhatnagar, 2008). Exosomes can carry RNA
such as mRNA, miRNA and genomic fragments, and a large component of proteins and
lipids among host cells leads to triggering of an anti-viral state in uninfected cells (Huang et
193
al., 2013; Schorey & Bhatnagar, 2008; Valadi et al., 2007). For example, HIV transfer of the
antiviral cytidine deaminase APOBEC3G via exosomes inhibits HIV replication in recipient
cells (Khatua et al., 2009).
Therefore virus-free supernatant from mock- or FCV-infected cells could be treated
with proteinase K and the ability to induce SG assembly monitored by immunofluorescence
microscopy to confirm whether a protein mediator is involved in the paracrine induction of
SGs. The presence of exosome particles in the supernatant should also be evaluated using
commercial exosome purification procedures and immunoblotting against the exosome
marker CD9 (Zeringer et al., 2015). Proteomics analysis from cell culture supernatant is
difficult due to the presence of highly abundant serum proteins. However supernatant could
be fractionated over a gel filtration column into several fractions, and the individual fractions
could be tested to determine whether they were able to induce SGs. These fractions could
then be analysed by SDS-PAGE and liquid chromatography–mass spectrometry (LC-MS).
Currently, this option is being investigated in collaboration with the proteomics facility at the
University of Cambridge (collaboration with Dr Kathryn Lilley).
Overall, the results from Chapter 4 suggest that a new mechanism of paracrine
signalling during viral infection is involved in the assembly of SGs in uninfected cells and it
is proposed that this unprecedented mechanism could play a key part in the antiviral
response.
194
Chapter 5
Regulation of P-Bodies formation
during
FCV and MNV-1 infection
195
5.1 Optimisation of the markers and conditions used for P-bodies detection
The previous results revealed that the assembly of SGs is impaired during FCV
infection. This prompted us to ask whether caliciviruses also have the ability to modulate the
formation of other cytoplasmic RNA granules in infected cells, namely PBs. To this end,
FCV and MNV-1 were used as surrogate model for human norovirus, in order to examine
their effect on PBs formation.
First, this work began with the optimisation of experimental conditions for
immunofluorescence detection of PB assembly. While PBs are constitutively form in the
cytoplasm of cells under normal conditions, their numbers and size increase upon stress
condition (Teixeira et al., 2005). Therefore, PBs markers optimisation was tested in
unstressed cells that support FCV or MNV-1 replication, CRFK and RAW264.7 cells
respectively, and monitored by immunofluorescence microscopy using Dcp1 and Xrn1 as
markers.
Among different concentrations of Dcp1 antibody tested, cytoplasmic foci were only
detected in unstressed CRFK and RAW264.7 cells at dilution of 1:50, while lower dilution
did not lead to the detection of PBs (Figure 5.1, 5.2 and data not shown). By contrast, no
signal could be detected for Xrn1, even when testing a wide range of dilutions from 1:50 to
1:500 (Figure 5.1, 5.2 and data not shown) using both CRFK and RAW264.7 cells.
196
Figure 5.1 Optimization of Dcp1 and Xrn1 antibody dilution in CRFK cells for
immunofluorescence studies. CRFK cells were fixed and stained for different
concentrations of Dcp1 (A-C) (red) and Xrn1 antibodies (D) antibodies as indicated,
followed by Alexa-555 secondary antibody. ToPro3 stained nuclei are shown in blue. Cells
were visualised using a ZEISS LSM META confocal laser microscope. The bar indicates a
size of 10 µm.
ToPro-3 Dcp1 Overlay
ToPro-3 Dcp1 Overlay
ToPro-3 Dcp1 Overlay
A
B
C
1:50 1:100 1:500
D 1:50
Xrn1 Overlay ToPro-3
197
Figure 5.2 Optimization of Dcp1 and Xrn1 antibody dilution in RAW264.7 cells for
immunofluorescence studies. RAW264.7 cells were fixed and stained for different
concentrations of Dcp1 (A) and Xrn1 (B and C) antibodies, as indicated, followed by
Alexa-555 secondary antibodies. ToPro3 stained nuclei are shown in blue. Cells were
visualised using a ZEISS LSM META confocal laser microscope. The bar indicates a size
of 10 µm.
1:50
1:100
Xrn1 Overlay ToPro-3
Xrn1 Overlay ToPro-3
B
C
ToPro-3 Dcp1 Overlay
1:50 A
198
As discussed for SGs in section 3.6, the concentration of foetal bovine serum (FBS)
can act as a source of stress on cells and affect the condition for infection. Therefore the
effect of FBS concentration on PB assembly was analysed by incubating CRFK cells with
different concentrations of FBS (0, 2 and 10%) and monitoring the impact on PBs by
immunofluorescence microscopy using Dcp1 as marker (Figure 5.3). Immunofluorescence
analysis revealed no significant differences in numbers of cells displaying PBs or the number
of PBs per cell using these different concentrations (Figure 5.3, and data not show).
Therefore 2% of FBS was chosen for all subsequent studies as we did with SGs.
199
Figure 5.3 Optimization of foetal bovine serum concentration for detect of P-bodies
in CRFK cells. CRFK cells were grown overnight using different concentrations of FBS
(A-C), cells were then fixed and stained for Dcp1 antibody (red; middle panels), followed
by Alexa-555 secondary antibody. ToPro3 stained nuclei are shown in blue. Cells were
visualised using a ZEISS LSM META confocal laser microscope. The bar indicates a size
of 10 µm.
0% FBS 2% FBS 10% FBS
ToPoro-3 Dcp1 Overlay
ToPoro-3 Dcp1 Overlay
ToPoro-3 Dcp1 Overlay
A
B
C
200
5.2 Effect of MNV-1 infection on P-bodies formation.
To investigate whether MNV-1 infection interfere with PBs formation, RAW264.7
cells were infected with MNV-1 at MOI of 30 for 18h, and examined by immunofluorescence
microscopy at 0, 4, 8, 12 and 18h post infection using Dcp1 as a marker for PBs, and NS7 as
marker for MNV-1 (Figure 5.4). Overall, a significant reduction in number of cells displaying
PBs was observed from 8 hpi onwards during MNV-1 infection relative to mock-infected
cells (Figure 5.4). This disruption was reflected with a drop in the number of cells displaying
PBs from 90.4 to 39.4% when compared to mock-infected cells at 8 hpi, 68.6 to 19.2% at 12
hpi, and 99.6 to 28.5% at 18 hpi (Figure 5.5). In summary these experiments suggest that
MNV-1 infection disrupt PB assembly during infection, and that this starts in the middle of
the replication cycle at a time where viral RNA synthesis begins. .
201
0 hpi
4 hpi
8 hpi
- MNV-1 + MNV-1
NS7 DCP1 Overlay
NS7 DCP1 Overlay
NS7 DCP1 Overlay
NS7 DCP1 Overlay
NS7 DCP1 Overlay
NS7 DCP1 Overlay
- MNV-1 + MNV-1
- MNV-1 + MNV-1
202
Figure 5.4 Detection of P-bodies during MNV-1 infection. RAW264.7 cells were infected
with MNV-1 at MOI of 30 for several time points as indicated, and then immunostained for
Dcp1 antibody (red) and NS7 (green) as markers for PBs and virus respectively. Left panels
show NS7 alone (green), middle panels show Dcp1 alone (red) and right panels show an
overlay of both. Cell nuclei were stained with ToPro-3 (blue). Cells were visualised using a
ZEISS LSM META confocal laser microscope. The bar indicates a size of 10 µm.
12 hpi
18 hpi
NS7 Dcp1 Overlay
NS7 Dcp1 Overlay
NS7 Dcp1 Overlay
NS7 Dcp1 Overlay
- MNV-1 + MNV-1
- MNV-1 + MNV-1
203
0 4 812
18
0
2 0
4 0
6 0
8 0
1 0 0
h o u rs p o s t- in fe c t io n
Ce
lls
dis
pla
yin
g
PB
s
(%)
- M N V -1
+ M N V -1n s
****
****
****
n s
Figure 5.5 Quantification of the number of cells displaying P-bodies during MNV-1
infection. RAW264.7 cells were infected with MNV-1 at MOI of 30 for several time points,
stained for Dcp1 and NS7 as described in Figure 5.4. Confocal images were then used to
quantify the numbers of cells with PBs using Image J software. Results shown are the
percentage of cells displaying PBs at each time point compared to the control, and are the
mean of three independent experiments (+/-SE; **** is P ≤0.0001; ns: not significant).
204
To confirm that this effect was not cell-line specific, BV-2 cells were infected with
MNV-1 at MOI of 30 for 12h and the effect on PB assembly was investigated using Dcp1 as
marker for PBs and NS7 as marker for the virus. Similarly to results obtained with
RAW264.7 cells, the infection with MNV-1 also resulted in the reduction of PBs in BV-2
cells (Figure 5.6). As shown on Figure 5.7, 27.8% of MNV-1 infected cells displayed PBs,
against 76.9.6% for mock-infected cells (Figure 5.7).
205
Figure 5.6 Detection of P-bodies during MNV-1 infection using BV-2 cells. BV-2 cells
were infected with MNV-1 at MOI of 30 for 12h, and then immunostained for Dcp1
antibody (red) and NS7 antibody (green) as markers for PBs and virus respectively. Left
panels show NS7 alone (green), middle panels show Dcp1 alone (red) and right panels show
an overlay of both. Cell nuclei were stained with ToPro-3 (blue). Cells were visualised using
a ZEISS LSM META confocal laser microscope. The bar indicates a size of 10 µm.
NS7 Dcp1 Overlay
NS7 Dcp1 Overlay
- MNV-1 + MNV-1
206
0
2 0
4 0
6 0
8 0
1 0 0
1 2 h o u rs p o s t- in fe c t io n
Ce
lls
dis
pla
yin
g
PB
s
(%)
- M N V -1
+ M N V -1
*
Figure 5.7 Quantification of the number of cells displaying P-bodies during MNV-1
infection in BV-2 cells. BV-2 cells were infected with MNV-1 at MOI of 30 for several
time points, stained for Dcp1 and NS7 antibodies as described in Figure 5.6. Confocal
images were then used to quantify the numbers of cells with PBs using Image J software.
Results shown are the percentage of cells displaying PBs compared to the control, and are
the mean of three independent experiments (+/-SE; * is P ≤0.5).
207
The section 5.1 revealed that using Xrn1 as marker did not allow the detection of PBs
in RAW264.7. However, we cannot exclude that this marker could accumulate in the PBs
formed in RAW264.7 cells during MNV-1 infection. To test this, RAW264.7 cells were
infected with MNV-1 at MOI of 30 for 24h and the assembly of PBs monitored by
immunofluorescence microscopy using Xrn1 and NS7 as marker for PBs and infection
respectively. As shown on Figure 5.8, no cytoplasmic foci could be detected using this
marker at any point during infection. These results confirm that Xrn1 is not a suitable marker
for the detection of PBs either during MNV-1 infection of RAW264.7 cells or during normal
conditions (Figure 5.2, Figure 5.8).
208
0hpi
4hpi
- MNV-1 + MNV-1
8hpi
NS7 Xrn1 Overlay
NS7 Xrn1 Overlay
NS7 Xrn1 Overlay
NS7 Xrn1 Overlay
NS7 Xrn1 Overlay
NS7 Xrn1 Overlay
- MNV-1 + MNV-1
- MNV-1 + MNV-1
209
Figure 5.8 Detection of P-bodies during MNV-1 infection using Xrn1 as a marker.
RAW264.7 cells were infected with MNV-1 at MOI of 30 for different time points, and then
immunostained for Xrn1 antibody (red) and NS7 (green) as markers for PBs and virus
respectively. Left panels show NS7 alone (green), middle panels show Xrn1 alone (red) and
right panels show an overlay of both. Cell nuclei were stained with ToPro-3 (blue). Cells
were visualised using a ZEISS LSM META confocal laser microscope. The bar indicates a
size of 10 µm.
12hpi
18hpi
24hpi
NS7 Xrn1 Overlay
NS7 Xrn1 Overlay
NS7 Xrn1 Overlay
NS7 Xrn1 Overlay
NS7 Xrn1 Overlay
NS7 Xrn1 Overlay
- MNV-1 + MNV-1
- MNV-1 + MNV-1
- MNV-1 + MNV-1
210
5.3 Effect of FCV infection on the assembly of P-bodies.
To investigate the effect of FCV infection on the formation of PBs, CRFK cells were
infected with FCV Urbana at MOI 0.2 for 6h, and the presence of PBs was investigated by
immunofluorescence microscopy using Dcp1 as marker for PBs and p76 as marker for viral
particles (Figure 5.9). As shown on figure 5.9, FCV resulted in a reduction in PBs abundance.
The quantification of the number of cells displaying PBs revealed that 91% of mock-infected
contained PBs, while only 19% of FCV-infected cells contained PBs (Figure 5.10). These
results suggest that FCV infection, like MNV-1 infection, disrupts PB assembly.
211
Figure 5.9 Detection of P-bodies during FCV infection. CRFK cells were infected with FCV at
MOI of 0.2 for 6h, and then immunostained for Dcp1 antibody (red) and p76 antibody (green) as
markers for PBs and virus respectively. Left panels show p76 alone (green), middle panels show
Dcp1 alone (red) and right panels show an overlay of both. Cell nuclei were stained with ToPro-3
(blue). Cells were visualised using a ZEISS LSM META confocal laser microscope. The bar
indicates a size of 10 µm.
p76 Dcp1 Overlay
p76 Dcp1 Overlay
-FCV
+FCV
212
0
2 0
4 0
6 0
8 0
1 0 0
Ce
lls
dis
pla
yin
g
PB
s
(%)
- F C V
+ F C V 6 h
***
Figure 5.10 Quantification of the number of cells displaying P-bodies during FCV
infection. CRFK cells were infected with FCV at MOI of 0.2 for 6h, stained for Dcp1 and
p76 as described in Figure 5.9. Confocal images were then used to quantify the numbers of
cells with PBs using Image J software. Results show are the percentage of cells displaying
SGs compared to the control, and are the mean of three independent experiments (+/-SE;
*** is P ≤0. 001).
213
5.4 Effect of chemical induction of oxidative stress on P-body assembly
PBs formation is induced by a wide range of experimental stresses such as glucose
deprivation, osmotic stress, and decreased translation initiation rates (Buchan & Parker, 2009;
Iwaki & Izawa, 2012; Teixeira et al., 2005). Importantly, most of these stresses are also
inducing the assembly of SGs (Anderson & Kedersha, 2006; Kedersha & Anderson, 2009;
Kedersha et al., 2005). By contrast, some inducer, such as Clotrimazol, can induce SGs but
not PBs (Buchan et al., 2011), and conversely, acidic stress only induces PBs formation
(Iwaki and Izawa, 2012). In addition, some studies have suggested that the assembly of stress
granules is dependent on the existence of PBs, implying that stress that lead to PB formation
could also induce SGs formation (Buchan et al., 2008). In contrast, other studies support
models in which mRNA exists in equilibrium between polysomes, SGs and PBs (Anderson &
Kedersha, 2009a; Decker & Parker, 2012; Kedersha et al., 2000; Lloyd, 2012) and (Figure
1.4). Finally, the size and numbers of PBs are dependent on the type of stress (Buchan &
Parker, 2009; Teixeira et al., 2005). As such, the induction of PBs by oxidative stress, such as
sodium arsenite or hydrogen peroxide, and the link with SG assembly, is a matter of debate.
Some studies demonstrated that PBs numbers and size increased upon sodium arsenite
treatment (Eulalio et al., 2007a; Eulalio et al., 2007b; Kedersha & Anderson, 2007; Serman
et al., 2007; Shah et al., 2013; Souquere et al., 2009; Wang et al., 2013; Wilczynska et al.,
2005). However, other studies demonstrated that hydrogen peroxide treatment results in the
inhibition of PBs, directly (Shah et al., 2013) or through the inhibition of factors, such as
microtubules, required for PB assembly, such as microtubules (Hu & Lu, 2014; Lee et al.,
2005; Lindsay & McCaffrey, 2011). Therefore, building on the previous results in section 3.2
214
and 3.8, suggesting that CRFK cells are able to form SGs in response to sodium arsenite or
hydrogen peroxide, the impact of these treatments on the assembly of PBs was investigated.
The ability of CRFK cells to respond to stresses that induce SGs assembly was
investigated by treating CRFK cells with 0.5 mM of sodium arsenite for 1h or with 1mM
H2O2 for 2h and subsequently monitoring the assembly of PBs by immunofluorescence
microscopy using Dcp1 as marker (Figure 5.11). Upon hydrogen peroxide treatment, the
number of PBs dropped from 95.2% to 8.8% (Figure 5.12). By contrast, a more modest yet
significant dispersion of PBs was observed upon treatment of cells with sodium arsenite with
75.2% of cell displaying PBs against 90.8% in mock-treated cells (Figure 5.12). These results
suggest that the ability of CRFK cells to form PBs is reduced in response to hydrogen
peroxide and sodium arsenite. Although only preliminary, this could reflect that the formation
of SGs is not strictly dependent on pre-existing PBs in CRFK cells. This would rather support
a model in which mRNAs are in equilibrium between RNA granules and whereby stimulating
the assembly of a type of granules, in this case SGs, conversely has a negative impact on the
abundance of other types, in this case PBs.
215
ToPro-3 Dcp1 Overlay
ToPro-3 Dcp1 Overlay
ToPro-3 Dcp1 Overlay
ToPro-3 Dcp1 Overlay
- H2O2
+ H2O2
- SA
+SA
Figure 5.11 Effect of the sodium arsenite and hydrogen peroxide treatment on P-
body assembly. CRFK cells were treated with 0.5mM sodium arsenite (SA) for 1h or 1
mM of hydrogen peroxide (H2O2) for 2h, stained for Dcp1 as a marker for PBs (red),
followed by Alexa-555 secondary antibody. Cell nuclei were stained with ToPro-3 (blue)
(left panels) and middle panels show Dcp1 alone (red) and right panels show an overlay of
both.
216
- S
A
+S
A
- H
2O
2
+H
2O
2
0
2 0
4 0
6 0
8 0
1 0 0
1 2 0
Ce
lls
dis
pla
yin
g
PB
s
(%)
*****
Figure 5.12 Quantification of the number of cells displaying P-bodies upon treatment
with the sodium arsenite and hydrogen peroxide. CRFK cells were treated with 0.5 mM
of sodium arsenite (SA) for 1h or 1mM hydrogen peroxide (H2O2) for 2h, stained for
Dcp1 as a marker for PBs, followed by Alexa-555 secondary antibody as describe in
figure 5.11. Confocal images were then used to quantify the numbers of cells with P-
bodies using Image J software. Results show the percentage of cells with PBs upon SA or
H2O2 compared to untreated cells, and are the mean of three independent experiments (+/-
SE; * and **** is p ≤ 0.1 and 0.0001 for SA and H2O2 respectively.
217
5.5 Discussion
5.5.1 Modulation of P-bodies assembly during MNV-1 and FCV infection
The translation and degradation of cytoplasmic mRNA is one of several mechanisms
used by eukaryotic cells in order to regulate gene expression (Cougot et al., 2004; Teixeira et
al., 2005). These mechanisms are thought to be competing pathways as the inhibition of
translation could result in an increased mRNA degradation rate (Franks & Lykke-Andersen,
2008). PBs play a role in the regulation of mRNA turnover and contain translationally
repressed mRNAs together with several proteins involved in mRNA decapping,
deadenylation, miRNA mediated mRNA silencing, and mRNA storage (Anderson &
Kedersha, 2009a; Cougot et al., 2004; Eulalio et al., 2007a; Eulalio et al., 2007b; Kedersha &
Anderson, 2009; Kedersha et al., 2005; Parker & Sheth, 2007; Teixeira et al., 2005). They are
found in cells under normal conditions and are conserved among eukaryotes (Decker &
Parker, 2012).
The results from chapter 3 showed that FCV infection impairs the ability of cells to
form SGs in response to different types of stresses. This could suggest that FCV infection has
the ability to prevent the inhibition of cellular protein synthesis, thereby enhancing viral
replication. While previous studies have identified that pre-existing PBs are require for SG
formation (Buchan et al., 2008; Kedersha et al., 2005), other did not (Mollet et al., 2008; Ohn
et al., 2008; Shah et al., 2013). It has also been suggested that the mRNA are in dynamic
equilibrium between polysomes, SGs and PBs (Balagopal & Parker, 2009; Buchan & Parker,
2009). These led to the hypothesis that caliciviruses may also interfere with cytoplasmic
RNA granules at the level of the PBs.
Thus to understand better the relationship between calicivirus infection and
cytoplasmic RNA granules, I investigated the regulation of PBs assembly during FCV and
218
MNV-1 infection. During MNV-1 infection, from 8 hpi, and FCV infection, from 6 hpi, PB
assembly is disrupted, as reflected by a reduction in Dcp1 cytoplasmic foci. MNV-1 infection
resulted in a reduction from 68.6 to 19.2% in the number of cells containing PBs, while FCV
infection resulted in a reduction from 90.8 to 19.1%. These results indicate that MNV-1 and
FCV have the ability to disrupt PBs, an event that may be required for a viral replication.
Viruses have evolved many strategies to counteract PB assembly and evade RNA
decay pathways. This disruption may be due to the modification or cleavage of a specific PB-
nucleating protein or may result from redistribution of PB components to enhance viral
replication (Lloyd, 2013; Reineke & Lloyd, 2013; Tsai & Lloyd, 2014). All these
mechanisms are dependent on the presence of specific viral products, RNA or proteins.
The disruption of PBs during HCV infection relies on an interaction between viral
proteins and PBs components (Ariumi et al., 2011), as well as an alteration of PBs
composition (Perez-Vilaro et al., 2014). HCV hijacks PBs component by redistributing
DDX6, Lsm1, Xrn1, PATL1, and Ago2 to the HCV replication factory localized around lipid
droplets, resulting in enhanced HCV replication. The flavivirus WNV can also co-opt
essential PB proteins, such as Ago2, DDX3, GW182, Xrn1 and Lsm1 for their replication by
re-localising these proteins to a viral replication centre (Chahar et al., 2013; Emara &
Brinton, 2007). An interaction between the viral proteins NS1 with the PBs component
RAP55 also results in inhibition of PB formation during influenza A virus infection (Mok et
al., 2012). Thus, to investigate the mechanisms by which FCV and MNV-1 infection disrupt
PBs, further studies should aim at dissecting the potential re-localization or association of
PBs components within caliciviruses replication complex using immunofluorescence
microscopy studies.
219
It has been shown that the picornavirus 3CPro can disrupt PB formation through the
cleavage of specific PB-nucleating proteins, such as Xrn1, Pan3, and Dcp1a (Dougherty et
al., 2011). We previously suggested that the inhibition of stress granules by FCV infection
might be due to the cleavage of the SG-nucleating protein, G3BP1 by the calicivirus 3CLpro
which has similar targets as the picornavirus 3Cpro (Sosnovtseva et al., 1999). We could
speculate that the repression of PB assembly by FCV and MNV-1 might result from the
cleavage of critical PB components, such as Dcp1. Further studies are therefore required to
investigate the cleavage of Dcp1 or other PBs components using immunoblotting of cell
lysates from FCV and MNV-1 infected cells. In addition, the overexpression of a wild type
FCV or MNV 3CLpro or an inactive mutant, and examination of the effect on PBs, should
indicate whether the calicivirus 3CLpro play a role in PB assembly.
In addition to these mechanisms, signalling pathways have also been linked with the
disruption of PBs. For example, the formation of PBs in response to glucose deprivation in
S.cerevisiae is dependent on the inactivation of the PKA signalling pathway (Ramachandran
et al., 2011). The activation of PKA signalling leads to the phosphorylation of Pat1, a critical
protein for PB assembly, impairing the interaction between Pat1 with other PB components,
such as the RNA helicase Dhh1, and inhibiting the PB assembly (Ramachandran et al., 2011).
As similar behaviour was observed under different stress condition, such as hyperosmotic
stress (Shah et al., 2013). By contrast, the activation of PKA has no effect on stress granule
formation (Shah et al., 2013). Previously, it has been reported that the activation of PKA is
necessary for porcine enteric calicivirus (PEC) replication (Chang et al., 2002). While other
PEC do not replicate in cell culture, the Cowden strain of PEC can replicate in a continuous
cell line (LLC-PK)-containing intestinal content (IC) from uninfected gnotobiotic pigs and
viral replication activates the PKA signalling pathway to inhibit IFN-mediated STAT1
activation (Chang et al., 2002; Chang et al., 2004). Therefore we could speculate that other
220
caliciviruses may also result in activation of PKA, in turns leading to Pat1 phosphorylation
and PBs disruption. To test this hypothesis, further work is required to investigate the
activation of PKA signalling and Pat1 phosphorylation during FCV and MNV-1 infection.
It has been demonstrated that the dissociation of Dcp1a from PBs is strongly
associated with activation of the JNK signalling pathway, part of the MAPK signalling
pathway, leading to the phosphorylation of Dcp1a at the residue S315 (Rzeczkowski et al.,
2011). Moreover, the localisation of DCP1a, Edc4, and Xrn1 to PBs is also affected by JNK
activation in human cells (Rzeczkowski et al., 2011). Previous work from our laboratory has
demonstrated that calicivirus replication is dependent on the activation of two of the main
arms of the MAPK signalling pathway, ERK and p38 (Royall et al., 2015). This activation is
critical to induce eIF4E phosphorylation and mediate host translational control during MNV-
1 infection (Royall et al., 2015). Thus, we could investigate whether calicivirus infection also
activate the remaining arm of the MAPK signalling pathway, namely JNK, using inhibitors of
JNK and immunoblotting studies.
5.5.2 Correlation between PBs and SGs assembly
The results from chapter 3 suggested that FCV infection impairs SGs assembly, while
this chapter revealed that FCV infection also impairs PBs formation. Given that some, but not
all, previous studies proposed that SGs assembly is dependent on pre-existing PBs (Buchan et
al., 2008), we could propose that the inhibition of SGs formation is a consequence of PBs
disruption. However the directional link between PBs and SGs is still controversial. Thus to
investigate the link between PBs and SGs in cells supporting FCV replication we studied the
impact of stress inducing SGs on PBs assembly. The results indicate that hydrogen peroxide,
221
which is potent inducer of SGs (Figure 3.21 & 3.22), disrupt PBs (Figure 5.12). Similarly,
sodium arsenite, a potent inducer of SGs (Figure 3.17 & 3.18), also disrupts PBs although to
a lesser extent (Figure 5.12). Therefore, these results would not support a model in which
SGs assembly is dependent on pre-existing PBs. In contrast, this would support the more
recent model in which mRNAs are found in dynamic equilibrium between polysomes and
cytoplasmic RNA granules (Anderson & Kedersha, 2009b; Kedersha & Anderson, 2009;
Kedersha et al., 2005). By inducing SGs formation with sodium arsenite or hydrogen
peroxide, this would displace the equilibrium in favour of SGs and have a negative knock-on
effect on PBs. However this results are only preliminary an further studies would be needed
to confirm this model, for instance by using stress that promote PBs and analysing the effect
on SGs assembly, or by monitoring the transition of marker mRNAs between the difference
RNA granules in response to stress.
Correlating with the potent effect of hydrogen peroxide on PBs disruption, several
studies have demonstrated an association between hydrogen peroxide treatment and
depolymerisation of microtubules, which is required to deliver mRNAs to PBs (Hu & Lu,
2014; Lindsay & McCaffrey, 2011). In addition, it was proposed that hydrogen peroxide also
plays a role in the phosphorylation of critical PB proteins components (Yoon et al., 2010).
Dcp2 protein is phosphorylated in response to hydrogen peroxide in Saccharomyces
cerevisiae, affect its association with other PB components and impairing their assembly
(Yoon et al., 2010). Furthermore, the hyper-phosphorylation of Dcp1 during cell mitosis also
results in PB inhibition (Aizer et al., 2013).
In conclusion the results from this chapter indicate that caliciviruses infection disrupt
PBs, and suggest several putative mechanisms responsible for this: via the cleavage of the
222
critical PB-nucleation protein, such as Dcp1, via an interaction between viral proteins and PB
components, through phosphorylation certain PB proteins, such as Dcp1 and Pat1 or via the
regulation of a specific signalling pathway, such as PKA or JNK pathways. The inhibition of
PB assembly upon oxidative stress that induce SG formation in the same cell line also lead to
suggest that these granules are formed independently from each other at least in our system.
223
Chapter 6
Conclusion
224
HuNoV is one of the most important causes of acute gastroenteritis worldwide and
responsible for approximately 50% of all gastrointestinal outbreaks, leading to more than
200,000 deaths annually. In the UK, more than a million people are infected every year and
the consequences of this illness amount to an economical loss of more than £100 million per
annum. Viruses of the Norovirus genus are members of the family Caliciviridae, subdivided
into six genogroups (GI-VI) containing several genotypes that infect human and animal hosts.
Due to the lack of suitable cell culture systems to study HuNoV, two animal caliciviruses,
FCV and MNV provide surrogate models to study norovirus biology and interactions with the
infected hosts. Using these models, previous studies have clearly highlighted that calicivirus
have evolved a complex relationship with their hosts to hijack to cellular translation
machinery and mediate viral translation.
In response to stresses such as viral infection, host cells have evolved several
responses leading to the temporary inhibition of protein synthesis to halt viral replication.
One of these responses is mediated by the formation of cytoplasmic RNA granules, SGs and
PBs, where mRNA are either translationally stalled or degraded, respectively. Moreover these
granules are thought to contribute to the immune response against viruses. Therefore, in order
to replicate, viruses have evolved different mechanisms to interfere the assembly of these
cytoplasmic granules. Thus, to better understand the relationship between caliciviruses and
the infected host, I have investigated the formation of RNA granules during caliciviruses
infection using FCV and MNV-1 as a model for HuNoV.
The results discussed in Chapter 3 suggest that infection with either MNV-1 or FCV
did not induce the formation of SGs. These observations however differ between MNV-1 and
FCV. Indeed, I could not detect the formation of SGs in RAW264.7 and BV-2 cells infected
225
with MNV-1 or mock cells that exposed to cellular stress such as sodium arsenite. The
absence of SGs in RAW264.7 and BV-2 cells may be attributed to the inability of these cells
to form stress granules. I propose that this effect may be due to the over-expression of the
protein MCPIP1 protein, which has previously been shown to impair SG formation in
RAW264.7 cells in response to stress or infection. Unlike RAW264.7 and BV-2 cells, CRFK
cells, that support FCV infection, have the ability to form SGs in response to cellular stress.
Furthermore, FCV infection prevents the assembly of SGs in CRFK cells, and actively
disrupts the assembly of SGs formed in response to cellular stresses such as sodium arsenite
and hydrogen peroxide. Moreover, these results suggest that this inhibition of SGs assembly
is associated with viral replication as UV-inactivated virus does not impact on SGs formation.
This implies the involvement of a yet unidentified viral product, RNA or protein, in the
inhibition of SGs response. Based on previous studies, I propose that the viral protease may
contribute to this by cleaving an important SG-nucleating protein or that non-structural
protein may sequester SG components. Further studies will aim at dissecting the molecular
basis of these observations.
Although FCV infection prevents SGs assembly in infected cells, some neighbouring
cells in the vicinity of infected cells displayed SGs, suggesting a cell-to-cell induction
mechanism via a biological or chemical signal explored in Chapter 4. Using virus-free
supernatant from FCV-infected cells, I demonstrated the paracrine induction of SGs
assembly. This is an unprecedented observation that could reflect a new antiviral stress
response. Our results show that this paracrine induction is not mediated by interferon or
soluble RNA molecule. In contrast, heat shock inactivated the supernatant ability to induce
SGs, suggesting that the signalling messenger could have a proteinaceous or more complex
origin.
226
Several studies have suggested that stress granules and perhaps P-bodies, may serve
as sentinel mechanisms for IFN activation by acting as a cytoplasmic platform that enhances
IFN and cytokine signalling through the condensation of IFN signalling moieties with their
ligands within RNA granules. Stress granules induced in a paracrine manner in uninfected
cells may therefore be important for priming neighbouring cells against infection.
Moreover this raises the question whether these granules may help us to develop
novel therapeutic or preventive strategy for viral infectious diseases. As this induction of SGs
prevent viral replication and triggers an antiviral response, the identification of the paracrine
SG inducer could lead to new strategies to block viral replication. As the mechanism targeted
is based on the aggregation of host proteins, this would be unlikely to lead to the
development of drug-resistant viruses. It also raises the possibility of developing novel and
broad antivirals therapeutics with broad spectrum activity.
Finally, in addition to the effect of infection on SGs, we have also analysed the
assembly of PBs during FCV and MNV-1 infection. The results discussed in Chapter 5
showed a strong disruption of PBs assembly during infection by either FCV or MNV-1.
Again, the association of the disruption of these granules with viral replication may also
reflect an important role for viral product in this disruption. This may occur via the cleavage
an essential PB-component by viral protease, an interaction of specific viral protein with
certain PB-component or phosphorylation of critical PB-components by activation a specific
signalling pathways could mediated this mechanisms of disruption. Further work will be
required to dissect the mechanisms leading to the PBs disruption during calicivirus infection.
227
In conclusion, Figure 6.1 summarizes the current working model of caliciviruses
interaction with the mRNA metabolism pathways and RNA granules. In this model, FCV
and MNV-1 infection prevent the inhibition of cellular translation in infected cells by
disrupting the assembly of SGs (FCV) or/and PBs (FCV and MNV). This may prevent the
induction of antiviral responses and maintain viral translation. On the other hand, the
paracrine induction of SGs in bystander cells may reflect a new antiviral pathway induced by
caliciviruses to control infection.
228
Figure 6.1 Proposed working model for the effect of FCV and MNV-1 on stress granules
and P-bodies. Left side shows that the infection prevent the inhibition of cellular translation
via disruption SGs (FCV) or PBs (FCV and MNV-1). Right side shows the paracrine
induction of SGs in bystander cells responding to FCV infection through an unknown
signalling molecule (?).
229
Chapter 7
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Appendix 1
0.1% Triton X-100
- 100 µl Triton X-100
- 100 ml MilliQ water
PBS/BSA
- 1g BSA (Sigma)
- 200 ml autoclaved PBS
4% Paraformaldehyde
- 8g Paraformaldehyde
- 200 ml autoclaved PBS
- 20 µl CaCl2
- 20 µl MgCl2
-
ToPro-3
- 1µl ToPro-3
- 10 ml PBS